An Experimental and Bioinformatics Protocol for RNA-seq Analyses of Photoperiodic Diapause in the Asian Tiger Mosquito, Aedes albopictus

Summary

RNA-Seq analyses are becoming increasingly important for identifying the molecular underpinnings of adaptive traits in non-model organisms. Here, a protocol to identify differentially expressed genes between diapause and non-diapause Aedes albopictus mosquitoes is described, from mosquito rearing, to RNA sequencing and bioinformatics analyses of RNA-Seq data.

Abstract

Photoperiodic diapause is an important adaptation that allows individuals to escape harsh seasonal environments via a series of physiological changes, most notably developmental arrest and reduced metabolism. Global gene expression profiling via RNA-Seq can provide important insights into the transcriptional mechanisms of photoperiodic diapause. The Asian tiger mosquito, Aedes albopictus, is an outstanding organism for studying the transcriptional bases of diapause due to its ease of rearing, easily induced diapause, and the genomic resources available. This manuscript presents a general experimental workflow for identifying diapause-induced transcriptional differences in A. albopictus. Rearing techniques, conditions necessary to induce diapause and non-diapause development, methods to estimate percent diapause in a population, and RNA extraction and integrity assessment for mosquitoes are documented. A workflow to process RNA-Seq data from Illumina sequencers culminates in a list of differentially expressed genes. The representative results demonstrate that this protocol can be used to effectively identify genes differentially regulated at the transcriptional level in A. albopictus due to photoperiodic differences. With modest adjustments, this workflow can be readily adapted to study the transcriptional bases of diapause or other important life history traits in other mosquitoes.

Introduction

Rapid advances in next-generation sequencing (NGS) technologies are providing exciting opportunities to probe the molecular underpinnings of a wide range of genetically complex ecological adaptations in a broad diversity of non-model organisms^1–3. This approach is extremely powerful because it establishes a basis for population and functional genomics studies of organisms with an especially interesting and/or well-described ecology or evolutionary history, as well as organisms of practical concern, such as agricultural pests and disease vectors. Thus, NGS technologies are leading to rapid advances in the fields of ecology and have the potential to address problems such as understanding the mechanistic bases of biological responses to rapid contemporary climate change⁴, the spread of invasive species⁵, and host-pathogen interactions^6,7.

The extraordinary potential of NGS technologies for addressing basic and applied questions in ecology and evolutionary biology is in part due to the fact that these approaches can be applied to any organism at a moderate cost that is feasible for most research laboratories. Furthermore, these approaches provide genome-wide information without the requirement of a priori genetic resources such as a microarray chip or complete genome sequence. Nevertheless, to maximize the productivity of NGS experiments requires careful consideration of experimental design including issues such as the developmental timing and tissue-specificity of RNA sampling. Furthermore, the technical skills required to analyze the massive amounts of data produced by these experiments, often up to several hundred million DNA sequence reads, has been a particular challenge and has limited the widespread implementation of NGS approaches.

Recent RNA-Seq studies on the transcriptional bases of diapause in the invasive and medically important mosquito Aedes albopictus provide a useful example of some of the experimental protocols that can be employed to successfully apply NGS technology to studying the molecular basis of a complex ecological adaptation in a non-model organism^8–10. A. albopictus is a highly invasive species that is native to Asia but has recently invaded North America, South America, Europe, and Africa^11,12. Like many temperate insects, temperate populations of A. albopictus survive through winter by entering a type of dormancy referred to as photoperiodic diapause. In A. albopictus, exposure of pupal and adult females to short (autumnal) day lengths leads to the production of diapause eggs in which embryological development is completed, but the pharate larva inside the chorion of the egg enters a developmental arrest that renders the egg refractory to hatching stimulus^15–17. Diapause eggs are more desiccation resistant^5,18 and contain more total lipids¹⁹ than non-diapause eggs. Photoperiodic diapause in A. albopictus is thus a maternally controlled, adaptive phenotypic plasticity that is essential for surviving the harsh conditions of winter in temperate environments. Despite the well-understood ecological significance of photoperiodic diapause in a wide range of insects^20,21, the molecular basis of this crucial adaptation is not well characterized in any insect²². In organisms such as A. albopictus that undergo an embryonic diapause at the pharate larval stage, it remains a particularly compelling challenge to understand how the photoperiodic signal received by the mother is passed to the offspring and persists through the course of embryonic development to cause arrest at the pharate larval stage.

This protocol describes mosquito rearing, experimental design and bioinformatics analyses for NGS experiments (transcriptome sequencing) performed to elucidate transcriptional components of photoperiodic diapause in A. albopictus. This protocol can be used for additional studies of diapause in A. albopictus, can be adapted to investigate diapause in other closely related species such as other aedine mosquitoes that undergo egg diapause²³, and is also more generally relevant to employing NGS approaches to study the transcriptional bases of any complex adaptation in any insect.

Protocol

1. Larval Rearing of Two A. albopictus Groups to Adulthood

1. Set two photoperiod cabinets with programmable lighting at 21 °C for optimal diapause expression¹⁶ and approximately 80% relative humidity.
   1. Program one cabinet for a 16L:8D light:dark cycle (a non-diapause inducing LD photoperiod). Set the second cabinet for an 8L:16D light:dark cycle (diapause inducing)¹³.
   2. Program ‘lights on’ at the same time in both cabinets to synchronize circadian time between photoperiods. 
2. Calculate the quantity of eggs needed to perform the experiment. Aim for 300-500 eggs per cage. At least three replicate diapause cages and three replicate non-diapause cages are needed for RNA generation. This totals to ca. 1,800-3,000 eggs per experiment.
   1. Hatch the eggs by submerging egg papers into ca. 500 ml of deionized H₂O.
   2. Add ca. 1 ml food slurry consisting of ground dog food and brine shrimp as previously described²⁴. Cover container with mesh, and keep the mesh in place with a rubber band.
   3. Place in the LD photoperiod cabinet for ca. 24 hr.
3. Transfer hatched larvae to 10 x 10 x 2 cm Petri dishes filled with ca. 90 ml deionized H₂O.
   1. Maintain ca. 30 larvae per dish. Transfer larvae to clean dishes every 48–72 hr, for example every Monday, Wednesday, and Friday (M-W-F)²⁴.
   2. Feed ca. 1 ml food slurry consisting of ground dog food and brine shrimp in deionized water every M-W-F as previously described²⁴. 
4. Set up three to four adult cages for each photoperiod treatment, where each cage comprises a biological replicate.
   1. From opposite sides of 9.5 L buckets, cut out one 10-by-14 cm hole, and another hole with 15 cm diameter. Cover the first with mesh. Cut approximately one foot length of an orthopedic stocking, and glue one end around the inside of the other hole.
   2. Cut the foot end of the stocking off and knot shut — open only when access to the interior of the cage is needed. For the cage lid, cut out all of the interior, leaving only the rim, and replace the interior plastic with mesh²⁴.
   3. Note the photoperiod, replicate number, cage start date, and other information relevant to the experiment with permanent marker on the side of the cage. 
5. Line the bottom of the adult cages with wet filter paper. Dampen the filter paper with enough deionized H₂O to increase local humidity in the cage, but avoid standing water, which can stimulate oviposition on the filter paper²⁴. Check the filter paper daily for drying, re-wet when necessary.
6. To produce sufficient eggs for an RNA library, include at least 100 females/9.5 L cage, with no more than 500 mosquitoes per cage.
7. Collect pupae M-W-F and place in a small cup of clean H₂O at a density of no more than 50 pupae per 25 ml H₂O. Transfer the pupae cup to an adult cage. Place cages in the respective photoperiod cabinet — A. albopictus pupae are photosensitive¹⁵.
8. Ensure daily that H₂O in cups is clean and clear, and remove dead pupae, because build-up of dead pupae can cause mass mortality. Remove H₂O cups after all pupae emerge.
9. Place organic raisins on the top mesh of the cage to provide sugar for emerged adults. Monitor raisins, and change them every 3-5 days to prevent mold accumulation.

2. Maintenance of Adults to Allow Mating and Egg Production

1. Maintain cages at high humidity (approx. 80%) by lining the cage bottom with a moist filter paper, and provide access to non-moldy raisins, as described above (Sections 1.5 and 1.9).

3. Blood-feeding

1. Prepare to blood-feed females between two to six days after eclosion to ensure that females have been exposed to at least eight unambiguous short days before oviposition commences for nearly 100% diapause eggs¹⁴.
2. Prepare the Hemotek Membrane Feeding system. Plug the feeding units into the power supply. Adjust the temperature of each unit to 37 °C using the adjustment screw. Use an electronic thermometer and probe to measure the temperature of the feeding unit during calibration.
3. Prepare the meal reservoir. Stretch a square of collagen feeding membrane over the aperture of the meal reservoir and secure it with an O-ring. Carefully pull the corners to remove wrinkles; trim the excess membrane with scissors.
   NOTE: Collagen membrane may not work well for all mosquito species, and it may be necessary to try several types to find the optimal membrane if working with a species other than A. albopictus. Parafilm works well with Culex pipiens.
   1. If blood is stored frozen, thaw at room temperature for at least 1 hr before using.
   2. Hold the reservoir so the membrane is facing down, unsupported, and the filling ports are facing up. Use a transfer pipette or syringe to fill the reservoir with approximately 3 ml of whole blood from chickens that has sodium citrate as an anti-coagulant. Seal filling ports with plastic plugs.
4. Attach the prepared reservoir to the feeder by screwing it onto the stud on the heat transfer plate on the bottom of the feeder. Invert the feeder and place it on top of the cage, membrane side down, so that mosquitoes can feed through the mesh of the cage. Keep the feeder on the cage for approximately 45 min to maximize feeding.

4. Stimulate Oviposition

1. Four to five days post blood meal, equip each cage with a dark colored 50 ml cup lined with unbleached seed germination paper (egg paper) or textured non-bleached paper towel and fill halfway with deionized water⁹. If more than 250 mosquitoes are in a cage, use two cups.
   NOTE: For small cages or single-female vials, “hay infusion” in oviposition containers may increase oviposition due to the odor of the microbial flora²⁵.

5. Collect and Store Eggs

1. Commence egg collection within 4-5 days of blood feeding because egg production typically peaks approximately five days after blood feeding, and then subsides over the next week.
2. Vary egg collection frequency on necessities of the experiment. For general purposes, collect egg papers on an M-W-F schedule. Remove egg papers from each cage and replace with fresh paper. Place recently removed papers in Petri dishes and store in SD photoperiod cabinet to avoid confounding effects of egg storage.
3. Allow egg papers to remain wet for 2 days post-oviposition to allow serosal cuticle formation, which increases egg desiccation resistance²⁶.
4. Approximately 48 hr post collection, dry eggs in open air. Dry the paper such that it is limp and slightly damp to the touch, but not so wet that the paper is dark from H₂O or stimulates hatching of eggs.
   NOTE: A 6.5” x 4” paper may take approximately 3.5 hr to dry. Be cautious not to over-dry egg papers, as this will result in egg desiccation²⁷.
5. Reserve additional eggs from both LD and SD photoperiods to assess diapause incidence and interpret the photoperiodic effect (see Measuring Diapause, Section 6).
6. For long-term storage, keep egg papers at 21 °C and approximately 80% humidity in Petri dishes. Keep Petri dishes in a Tupperware storage container with a flask of water to maintain local humidity as embryonic development takes four to five days at 21 °C.

6. Measure Diapause Incidence

1. Use additional reserved embryos (see Stimulate Oviposition, Section 4) that are 7–20 days old to quantify the diapause response.
2. Record the number of eggs present on each egg paper.
3. Stimulate eggs to hatch by completely submerging individual egg papers in a 90 ml Petri dish with approximately 80 ml deionized H₂O. Add approximately 0.25 ml food slurry.
4. After 24 hr tally the number of hatched first instar larvae. Place the Petri dish on a black surface to visualize larvae and place a light source on one side of dish. Larvae will move away from the light source, allowing for a clear tally of individual larvae. Remove individual larvae with a pipette while counting to prevent recounting individual larvae.
5. Place egg papers in a new Petri dish and re-dry. Re-hatch eggs after ~1 week and again tally eggs hatched using the above method.
6. Place egg papers with the remaining un-hatched eggs in new 90 ml Petri dishes with approximately 80 ml bleaching solution²⁸. Ensure that the egg papers are completely submerged in the bleaching solution and leave under a fume hood overnight to avoid the odor of bleach.
   NOTE: Bleaching solution can be stored for ~1 week at 4 °C, but should otherwise be made fresh.
7. Inspect eggs using a light microscope as the bleaching will clear the chorion and allow visualization of embryonated, un-hatched eggs. If the egg is embryonated, the egg will have an off-white color with eyes appearing as two small black dots opposite each other on the dorsal side. Tally the number of un-hatched, embryonated eggs¹³.
8. Determine diapause incidence with the following formula: % diapause = no. embryonated un-hatched eggs / (no. hatched eggs + no. embryonated un-hatched eggs) x 100 ¹³.

7. RNA Extraction from Eggs/pharate Larvae

NOTE: Use Trizol in a laminar flow hood.

1. Brush mosquito eggs containing developing embryos or pharate larvae at distinct development time points from egg papers to glass grinders using a camel-hair brush. Grind the eggs in Trizol (1 ml per 50–100 mg of tissue) until completely pulverized. Use at least 400 eggs per library to yield sufficient RNA.
   1. Alternatively, snap freeze eggs in liquid nitrogen and stored at -80 °C in microcentrifuge tubes for up to a month before grinding in Trizol.
2. Perform RNA extraction in Trizol followed by isopropanol precipitation according to manufacturer’s instructions.
3. Treat the bench with RNase decontamination solution or other agents to remove any residual nucleases to avoid RNA degradation.
   1. Treat the extracted RNA with DNase. According to manufacturer’s instructions, incubate the RNA samples with DNase for 30 min at 37 °C. Use 1 µl DNase for up to 10 µg of RNA in a 50 µl reaction. Increase the amount of DNase if there are more than 10 µg of RNA in one reaction.
   2. Inactivate DNase by adding 5 µl suspended DNase inactivation reagent. Incubate 5 min at room temperature, mixing three times during incubation period (gentle vortexing).
   3. Centrifuge at 10,000 x g for 1.5 min. Transfer the supernatant containing the treated RNA samples to fresh tubes for subsequent steps.
4. Assess the quality of the total RNA samples by fluorometry. Send the samples to a specialty facility with proper instrument for this task. The facility will perform on-chip gel electrophoresis to determine the sizes of RNA species in the sample, visualized by fluorescent dye instilled in the chip. The results will be returned as an electropherogram.
   1. Determine the integrity of total RNA samples by the presence or absence of degradation products, as evidenced by the presence of peaks between the 18S and 5S ribosomal RNA peaks on the resulting electropherogram (Figure 1B).

8. RNA Sequencing

1. Send total RNA samples with sufficiently high quality (Figure 1A) and quantity (usually >3 μg per library) to a commercial sequencing center for construction of enriched paired-end mRNA libraries and mRNA sequencing, following standard protocols.
2. If more than one lane is being used for sequencing a single experiment, split individual libraries into two lanes for sequencing to account for technical variation among lanes during sequencing.

9. lllumina Read Cleaning

NOTE: Figure 2 summarizes the bioinformatics portion of this protocol. For a full list of all programs and resources used in the bioinformatics section of this protocol, refer to Table 1. In addition, Supplemental File 1 contains command line examples for each of the following bioinformatics protocol steps.

1. Use ssaha2²⁹ (Table 1) to identify matches of 95% identity or higher to the NCBI UniVec Core database (Table 1), A. albopictus rRNA sequence (GenBank #L22060.1), and sequencing adapters (detailed command-line examples provided in Supplemental File 1). Remove read pairs with matches using Perl or a similar scripting tool, for example by adapting the provided Perl script (Supplemental File 2).
2. Clean remaining reads with the SolexaQA package³⁰ (Table 1; Supplemental File 1): trim regions with a phred score equivalent of less than 20 using the default settings of DynamicTrim.pl.
   1. Remove reads shorter than 25 bp with LengthSort.pl on both forward and reverse reads simultaneously. Evaluate the quality of the cleaned fastq files with FastQC (Table 1) — in particular, verify that the per-base sequence quality and the per-sequence quality scores are above 20.

10. Digital Normalization

1. Perform one round of digital normalization on the cleaned reads using the khmer tool³¹ (Table 1; Supplemental File 1), specifically normalize-by-median.py (using k-mer size 20, a coverage cut-off of 20, and x = 1e10).
2. Alternatively, if a machine with high RAM is available (hundreds of GBs), use Trinity's normalize_by_kmer_coverage.pl script (Table 1).

11. De Novo Transcriptome Assembly

1. Obtain access to a computer or computer cluster with up to 256 Gb of RAM and 24 cpus, depending on the size of the assembly.
2. Use Trinity³² (Table 1; Supplemental File 1) to assemble the digitally normalized read set into contigs. To reduce memory usage, use --min_kmer_cov 2.

12. Assembly Evaluation

1. Run assemblathon_stats.pl from the Assemblathon2 project³³ on the Trinity contig output. This script performs basic calculations relevant to evaluating assembly quality, such as number of scaffolds, N50, assembly composition, and more (Table 1; Supplemental File 1).

13. Annotation of the Assembled Transcriptome

1. Perform Blastx (Table 1) of the assembly against a reference protein set; for mosquitoes, Drosophila melanogaster, Anopheles gambiae, Culex pipiens, and Aedes aegypti are suitable references. Specifically, format the reference protein fasta file for blast, followed by blastx (Supplemental File 1).

14. Map Reads to the Assembly Using RSEM³⁴ (Table 1)

1. Create a 'transcript-to-gene-map' file, in which the first column contains the reference gene IDs, and the second column the contig IDs. In a spreadsheet editor, swap the first and second columns from the Blastx output, and write these columns to a .txt file. Use the LineBreak program to convert line breaks in the resulting .txt file to Unix format.
2. Create a reference dataset from the transcriptome fasta file using the rsem-prepare-reference script, provided in the RSEM package (Supplemental File 1).
3. Calculate the expression values separately for each library using the rsem-calculate-expression command, provided in the RSEM package (Supplemental File 1). As reads, use the paired fastq files resulting from the read-cleaning step (step 9.2).
   1. If RNA from a biological replicate was split into two lanes for sequencing, include both fastq files in the expression calculation to generate a single file.
4. Convert the expression results from each library to a matrix easily processed by other programs using the provided script rsem-generate-data-matrix, provided in the RSEM package (Supplemental File 1).

15. Differential Expression Analysis

1. Install R and EdgeR (Table 1).
2. Use read.delim to load the RSEM results from step 14.3 (Supplemental File 1). If necessary, round the counts to the nearest integer.
   NOTE: The EdgeR guide recommends limiting the dataset to genes with high enough expression to detect significance.
3. To format the data for EdgeR, generate a DGEList object from the loaded data file (Supplemental File 1). Then, normalize the data using TMM normalization (Supplemental File 1). Estimate the common and tagwise dispersions of the data (Supplemental File 1).
4. Identify differentially expressed genes with a Benjamini-Hochberg corrected p-value <0.05 (Supplemental File 1). Plot the distribution of log-fold-change vs. abundance (Supplemental File 1).

Results

Fluorometry of two representative RNA samples showed two bands at approximately 2,000 nt (Figure 1A, B). The insect 28S ribosomal RNA is comprised of two polynucleotide chains held together by hydrogen bonds, which are easily disrupted by brief heating or agents that disrupt hydrogen bonds³⁵. The resulting two components are approximately the same size as the 18S ribosomal RNA. The second RNA sample showed high levels of degradation (Figure 1B).

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Discussion

This protocol presents methods to discover differentially expressed genes due to photoperiodically induced diapause in A. albopictus. The protocol is significant in that it uniquely combines mosquito rearing and bioinformatics techniques to make all experimental aspects of a molecular physiology program accessible to novice users — in particular for those focusing on the photoperiodic diapause response. Existing methods, to our knowledge, do not provide as much detail in the rearing protocol — which ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Incubator - Model 818	Thermo-Scientific	3751	120 V
Controlled environment room	Thermax Scientific	N/A	Walk-in controlled environment room built to custom specifications by Thermax Scientific Products. A larger alternative to an incubator. http://thermmax.com/
Cool Fluorescent bulb	Philips	392183	4 W
Petri Dish 100 mm x 20 mm	Fisher	08-772-E
Filter Paper 20.5 cm	Fisher	09-803-6J
9.5 L Bucket	Plastican	Bway Products	http://www.bwayproducts.com/sites/portal/plastic-products/plastic-open-head-pails/117
Utility Fabric-Mosquito Netting White	Joann	10173292	http://www.joann.com/utility-fabric-mosquito-netting-white/10173292.html
Orthopedic stockings	Albahealth	23650-040	product no. 081420
Organic Raisins	Newman's Own	UPC: 884284040255
Oviposition cups (brown)	Fisher Scientific	03-007-52	The product is actually an amber 125 ml bottle that we saw the top off of.
Recycled Paper Towels	Seventh Generation	30BPT120
Modular Mates Square Tupperware Set	Tupperware		http://order.tupperware.com/pls/htprod_www/coe$www.add_items
Glass Grinder	Corning Incorporated	7727-2	These Tenbroeck tissue grinders break the eggs and release RNA into the TRI Reagent.
TRI Reagent	Sigma Aldrich	T9424	Apply 1 ml TRI Reagent per 50-100 mg of tissue. Caution — this reagent is toxic.
TURBO DNA-free	Ambion/Life Technologies	AM1907	This kit generates greater yield than traditional DNase treatment followed by phenol/chloroform cleanup, and it is simpler to use.
RNaseZap	Ambion/Life Technologies	AM9782	Apply liberally on the bench surfaces and any equipment that might be in contact with the RNA samples. The solution is slightly alkaline/corrosive, can cause irritation and is harmful when swallowed.
2100 Bioanalyzer	Agilent Technologies	G2939AA	Place up to 12 RNA samples on one chip.
Hemotek Membrane Feeder	Hemotek	5W1	This system  provides 5 feeding stations that can be used simultaneously. Includes PS5 Power Unit and Power cord; 5 FUI Feeders + Meal Reservoirs and O-rings; Plastic Plugs, Hemotek collagen feeding membrane; Temperature setting tool; and Plug extracting tool. The company's mailing address is: Hemotek Ltd; Unit 5 Union Court; Alan Ramsbottom Way; Great Harwood; Lancashire, UK; BB6 7FD; tel: +44 1254 889 307.
Digital Thermometer and Probe	Hemotek	MT3KFU	MicroT3 thermometer and KFU probe. This is used to set the temperature of each FUI feeding unit.
Chicken Whole Blood, non-sterile with Sodium Citrate	Pel-Freez Biologicals	33130-1	The 500 ml of blood were frozen and stored in 20 ml aliquots at -80 °C for up to 1 year. Thaw blood at room temperature for at least 1 hr before using.