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Of course we can! So, where do we start? Well, this project involves designing fusion proteins for endocytic assay development in murine cell lines so we will do everything in the genetic background of the domestic house mouse, Mus musculs, which is important for codon optimization skewed toward mammalian-specific tRNAs. This company wants to create a Cro DNA-binding domain fused to a aldehyde dehydrogenase expressed in the yeast Saccharomyces cerevisiae, and it wants to use that protein in a proprietary enzymatic kit. Our target protein can easily be found searching through the National Center for Biotechnology Information’s (NCBI’s) online search tool, which accesses publicly available databases, including the standard non-redundant database and additional rRNA and ITS databases. The proper name of the target enzyme in humans is aldehyde dehydrogenase 2 family member. A quick BLAST of the protein against the murine genetic database gives us our target sequence. In mice the official full name of the gene is aldehyde dehydrogenase 2, mitochondrial, and it is positioned on chromosome 5. It has 12 exons and the coding sequence is 1,560 base pairs. At this point, we can codon optimize the gene for expression in S. cerevisiae, which can be done using a simple python script. Because nucleotide triplets can present more combinations than there are amino acids and associated tRNA, there is redundancy in the genetic code, and several different 3-nucleotide codons can express the same amino acid. This codon bias is what necessitates codon optimization, and by looking at the coding sequences of the tRNA repertoire of your target organism, you can interchange codons based on your expression system. At this point, we’ve accumulated a lot of genetic data, and workflows with detailed protocols and images aid in collaboration and management of downstream experiments. We can organize all of these for you and present them in a format suitable for presentation at conferences and lab meetings.

Verifying your primers in silico is important to ensure that all the pieces will fit together in the final synthesis, and that terminal vectors will be properly expressed and presented in the biological context for which they are intended. The opensource, free website, Benchling, is a great place to input your template and primer sequences for amplicon assembly and downstream cloning reactions. Once your sure that the primers are going to work, you can order them from a reliable vendor such as, Integrated DNA Technologies (IDT) or EuroFins. They shouldn’t cost more than $10/primer pair, depending on the size and composition. Primers will usually be shipped to you at room temperature, in an envelope as lyophilized tubes. Always spin the 2 ml tubes in a microcentrifuge prior to reconstitution in water. Once they are spun, carefully add nuclease-free water to a 100 µM final concentration. It is usually around 100 - 500 µls. Once your primers are in solution, make sure to keep them on ice. Vortex and spin before storing at -20°C. These are your primer stocks. If for whatever reason your primers begin to lose their ability to amplify your desired product, always make a new working mix from your stock primers. The concentration of the working stock should be 10 µM, so about 1 µL will be added to a 25 µL PCR reaction. Once your primers are prepped, you only need to obtain some clean template DNA for amplification. Most PCR kits require ~50-250 ng of genomic DNA or 1 pg-10 ng of plasmid DNA. Plasmid DNA is circular and significantly less contaminated with off-target binding-sites because of it’s small size relative to chromosomal DNA.

INTRODUCTION

            Since before recorded history, humans have been plagued by arthropods as harbingers of disease and discomfort. Diseases, transmitted by arthropod vectors account for more than 700,000 deaths per year, more than 17% of all infectious disease (WHO, 2020). As a vector of the highly pathogenic Plasmodium parasite, the causative agent of malaria, mosquitoes have shaped the course of human history. Additionally, they are responsible for the transmission of Zika virus, yellow fever virus, Chikungunya virus and dengue to name a few and their persistence, geographic ubiquity and zoonosis make them a concern as urbanization and climate change usher in an uncertain future (Kraemer et al., 2019). Bed bugs are believed to have evolved to human blood feeding during their cohabitation in caves, where they made the jump from bat to human host (Sailer, 1952). Bed bugs do not represent a commensurate public health concern, but they do offer insight into a model blood feeding arthropod with a unique molecular profile. While modern society offers new solutions to old problems with an ever-expanding array of molecular tools, we are still burdened by vector-borne disease and pest infestations. Only through specifically addressing each challenge directly can these molecular methods begin to make an impact on arthropods as a public health concern.

The resurgence of C. lectularius

            In the United States, bed bug infestations were greatly reduced by the 1950s. Since then, C. lectularius has had a major resurgence in the US and across the world (Doggett, Geary and Russell, 2004; Potter, 2004, 2006) and prior to World War II, it was estimated that up to a third of European cities were plagued by bed bug infestations (Davies, Field and Williamson, 2012). The re-emergence occurred because of the development of dichlorodiphenyltrichloroethane (DDT) resistance (Anderson and Leffler, 2008), urbanization (Hwang et al., 2005), and global connectivity (Hentley et al., 2017) to name a few. The medical importance is largely due to allergy (Goddard and DeShazo, 2009) but infestations can cause psychological sequelae as well, including anxiety, insomnia and avoidance behavior (Goddard and De Shazo, 2012). There is also a significant economic burden associated with bed bugs as landlords are reluctant to disclose infestations to incoming tenants and old tenants carry the arthropods to new residence in their clothes and belongings (Scarpino and Althouse, 2019). While modern methods have been able to characterize and define why bed bugs are so persistent there have been few effective strategies developed that can combat infestations and bed bugs remain a serious nuisance pest.

The biology and genetics C. lectularius

            The most important environmental factor influencing survival and development in C. lectularius is temperature with the threshold for nymphal development, egg hatching and adult activity being between 13°C and 15°C (Usinger, 1966). Temperature extremes tolerated by bed bugs for short periods range from -15°C to 45°C (Omori, 1941). At 30°C, the most rapid developmental time from egg to adult is achieved in 25 days (Omori, 1941). The bed bug goes through 5 nymphal stages before becoming a sexual mature adult and each nymphal stage requires a blood meal to develop to the next stage. Adults perform a unique sexual behavior known as traumatic insemination in which the male penetrates the female’s abdomen through an external organ called the ectospermalage. Females can alter their immune physiology in anticipation of traumatic insemination because of the behaviors link to blood feeding (Siva-Jothy et al., 2019). At 23°C and 75% relative humidity, unfed females had an average weight of 4.98 mg and took in a 7.6 mg blood from which they could produce 8.87 eggs on average (Usinger, 1966). A mated female will feed for about 5 to 10 min after which she will locate a harborage to digest and develop eggs. At 27°C, young adults are stimulated to search for another blood meal every 3 days and if meals a readily available, egg laying can be continuous, with a maximum number of eggs laid by a single female found to be 541 (Titschack, 1930). Bed bugs cannot detect a host beyond 5 feet, meaning they have to be in close proximity for sustained periods to obtain a blood meal (Usinger, 1966). In C. lectularius host preference is for humans, although one study found that during cohabitation with cats and mice, bed bug samples were found to have feline blood in their midguts (Potts et al., 2021).

            The modern era of genomics has brought with it a new understanding of C. lectularius with the genome offering the potential to develop new molecular methods of analysis (Jeffrey A. Rosenfeld et al., 2016). Using both Pacific Biosciences and Illumina sequencing technologies, the genome of C. lectularius was found to be 697.9 Mb on an assembly with an N50 of 971 kb (Jeffrey A. Rosenfeld et al., 2016). They found a total of 36,985 coding and non-coding genes and developed RNA-seq transcriptomes for each developmental stage. There was a significant change in gene expression during blood feeding, including a number of genes expressed from the endosymbiont, Wolbachia, which represents a unique metabolic adaptation found in hematophagous arthropods (Jeffrey A. Rosenfeld et al., 2016). Development from the 2nd to the 5th nymphal stage was associated with the lowest amount of differential gene expression, with a large increase during sexual development from 5th stage nymphs to either male or female adults. The range of genes expressed throughout the lifecycle of C. lectularius ranged from 14,752 – 20,673 genes at any one time detected at levels above one read per kilobase per million reads (RPKM).

Diversity amongst invertebrate salivary genes and proteins

            Bed bugs belong to one of two sub-orders of Hemiptera, the Heteroptera, which comprises over 40,000 members from 89 families and includes phytophagous, predaceous, and hematophagous species (Walker et al., 2016). This contrasts with the other order of Hemiptera, the Homoptera, which is comprised of insects that feed exclusively on plant liquids (Krinsky, 2019). Hematophagous behavior evolved from the diversification of the phytophagous Heteropterans and was driven by the adaptation of a venom transmitting apparatus from piercing and sucking mouthparts (Cohen, 1995). Predaceous and hematophagous heteropteran venoms contain disulfide-rich peptides, bioactive molecules, and toxic enzymes but a comprehensive analysis of all salivary and venom components from most species is lacking (Walker et al., 2016).

            A recent study characterized the salivary transcriptomes of 9 different Heteropteran insects and contrasted profiles between phytophagous, predaceous, and hematophagous species (Yoon et al., 2021). Species representatives were included from predaceous (Epidaus tuberosus, Himacerus apterus, Sphedanolestes impressicollis, Laccotrephes japonensis, and Muljarus japonicus), hematophagous (C. lectularius), and phytophagous (Anoplocnemis dallasi, Graphosoma rubrolineatum, and Hygia opaca) Heteropteran samples. Genes expressed at high levels included digestive enzymes (cathepsin and serine protease), an anti-inflammatory protein (cystatin), a paralytic neurotoxin (arginine kinase), hexamerin, and an odorant binding protein. Hematophagous and predaceous species had increased transcription of salivary genes involved in cytolysis and proteolysis, unlike the phytophagous heteropterans, who had increased expression levels of vitellogenin, a multifunctional allergen and no expression of cytolysis and proteolysis genes (Yoon et al., 2021). Surprisingly, the salivary transcriptomic profile of C. lectularius was most similar to the phylogenetically distant Japanese water scorpion, Laccotrephes japonensis, who has a distinct feeding strategy although they both feed on vertebrates. They found S1 proteases with complement C1r/C1s, Uegf, Bmp1 (CUB) domains and clear secretory signals at significantly high levels. S1 proteases are serine proteases that have been shown to elicit IgE allergic responses and can act as allergens. They found odorant binding protein transcripts in high amounts across all heteropterans, which is consistent with known levels in C. lectularius, approaching nearly 40% of all salivary proteins (Francischetti et al., 2010a), and underscores their importance in detecting and sensing prey. Because C. lectularius doesn’t share a common ancestor with other hematophagous Heteropterans, it expresses novel proteins in abundance, including the acetyl/butyryl cholinesterase and Nudix-type hydrolase genes (Yoon et al., 2021).

            Looking at similarities and differences in the evolution of hematophagous insect genes can give insight into important functionalities within the salivary gland as well as other important tissues. Another study looked at convergent evolution across different parameters within hematophagous insect lineages (Freitas and Nery, 2020). They noted that evolutionary convergence has been demonstrated at the molecular level through amino acid substitutions (Foote et al., 2015) rates of mutation (Partha et al., 2019) and gene family copy number (Griesmann et al., 2018), and wanted to explore these possibilities computationally in hematophagous insects. Recent bioinformatic advancements have facilitated the development of computational tools to compare genetic data across many disparate lineages (Sackton and Clark, 2019), which is useful when analyzing gene family expansion amongst hematophagous insects that require high levels of enzymatic power to utilize the chemical energy from a blood meal (Nyanjom et al., 2018). They looked at genomic data from six independently evolved hematophagous lineages, Culicidae, Psychodidae, Glossinidae (Diptera order), Reduviidae, Cimicidae (Hemiptera order) and Phthiraptera (Psocodea order). They found that the HSP20 (EOG090W0E1I) gene family expanded rapidly and independently in both Culicidae and Cimicidae and represented the only true example of convergent evolution where the expansion happened after the transition to blood feeding (Freitas and Nery, 2020). This is consistent with previously published work demonstrating increased expression levels of heat shock protein genes after blood feeding in mosquitoes, kissing bugs and bed bugs, which may protect from the heat stress caused by blood feeding (Benoit et al., 2011; Paim et al., 2016).

The nitrophorins of C. lectularius

            Blood feeding behavior has evolved independently many times in invertebrates, creating unique molecular protein repertoires that accomplish the same aim of obtaining and metabolizing a blood meal. In proteins, selective pressures often drive the evolution of common functionalities in disparate structural starting points. For example, bed bugs and kissing bugs both have nitrophorin proteins that release nitric oxide (NO) in the host bloodstream, but the proteins are structurally unique and work by a different biochemical mechanism. Another example, in mosquitoes, ticks and kissing bugs, the biogenic amine scavenger lipocalin and D7 proteins represent a common functionality developing through convergent evolution among three disparate groups of organisms (Calvo et al., 2006). The independent evolution of the same functionality over millions of years from different genetic origins may signal nitrophorin’s physiological importance.

            In higher vertebrates, NO acts to regulate blood pressure, heal wounds, form memories and fight infections (Gantner, LaFond and Bonini, 2020). Nitric oxide is highly reactive and is produced in low quantities by NO synthase through the conversion of l-arginine and oxygen to citrulline and NO (Alderton, Cooper and Knowles, 2001). Nitric oxide is a free-radical with a half-life of under 1 second in biological tissues, requiring the need for a molecular chaperone to deliver it to its destination. The S-nitroso (SNO) reaction side product formed with free thiol groups in cysteine residues stabilizes NO in heme proteins, is reversible and regulates NO storage in the salivary gland of C. lectularius. The salivary glands of most hematophagous insects are acidic, preventing the nucleophilic attack of the nitrophorin-SNO complex. During a blood meal, C. lectularius delivers nitrophorin-SNO to the host where a higher pH and concentration gradient results in the release of free NO, which acts as a vasodilator in this context, driving blood flow to the site of the bite (Ribeiro et al., 1993). The nitrophorin heme is in the ferric (FeIII) redox state, which has no affinity to oxygen and a 7-fold reduced affinity to NO when compared to the ferrous heme of globin molecules. The mechanism of NO binding in the nitrophorin of C. lectularius was elucidated in 2004 (Weichsel et al., 2005a). At that time, the genome of C. lectularius was undetermined and the copy number of the nitrophorin gene was unknown, meaning that the most abundant nitrophorin in the salivary gland, as defined by an expressed sequence tag (EST), was used as a representative (Valenzuela and Ribeiro, 1998). The sequence of that EST was expressed from the pET17b expression vector in BL21DE3 Escherichia coli and the protein was conjugated to heme prior to x-ray crystallographic analysis down to a resolution of 1.75-Å by using both MAD and multiple isomorphous replacement anomalous scattering phasing (Weichsel et al., 2005a). This 32-kDa nitrophorin protein, designated cNP, is structurally unique to the nitrophorins of Rhodnius prolixus, but shows similarities to the functionally unrelated exonuclease (Mol et al., 1995) and inositol polyphosphate 5-phosphatase (Tsujishita et al., 2001) genes. The heme is contained within a hydrophobic fold made up of a repeated beta-sandwich motif, and is bound to a Cys-60 (2.42 Å) within a proximal alpha-helix (Weichsel et al., 2005a). The thiolate anion of the Cys-60 is stabilized by the N-terminus of the proximal helix and is hydrogen bound to Gln-56, further stabilizing NO-binding (Poulos, 1996). The cNP distal pocket has a 35-Å3 volume and carries 3 solvent molecules that assist in stabilization of the heme iron (Weichsel et al., 2005a). The formation of the SNO conjugate results in a conformational change within the cNP pocket with the Fe-N-O conjugate displaying a 119° bend and a bond length of 1.86 Å between Fe and N, indicating that the Cys-60 may still be bound to the heme iron. The Cys-SNO conjugate lies within a hydrophobic pocket bounded by Phe-64, Ala-21 and heme and is highly mobile with a mobility range of 50-Å2 (Weichsel et al., 2005a). Using static and stopped-flow electronic absorption spectroscopies, the mechanism and order of NO-binding dynamics to cNP was determined. Adding cNP to 2 mM NO at pH 5.6 produced a Soret banding trace indicated sequential binding of NO to ferric heme and coordination to the thiolate of Cys-60, followed by the binding of an additional NO to the thiolate anion and the transition to ferrous Fe. These findings defined the reversible mechanism of NO binding in C. lectularius, specifically that the Fe in cNP oxidizes S- and NO to Cys-SNO, with the subsequent binding of a second NO to the Cys-60, resulting in electron abstraction by Fe and the neutralization of the SNO conjugate (Weichsel et al., 2005b). This process is simply reversed as nitrophorin enters the host blood where pH levels increase, and NO concentrations are negligible.

Blood feeding behavior in mosquitos

            For hematophagous arthropods, finding a host is essential, and when it comes to vector-borne pathogens, the vector bite is the most critical component of disease transmission in humans, and mathematical models often assign host-vector contact rate as the most important risk factor for predicting disease risk (Ellis et al., 2011; Gao et al., 2016).        In bed bugs, finding a host means predominantly sensing non-visual cues like CO2, odor, and heat (Aak et al., 2014). These sensations require a host of molecules that react to physical, chemical, or optical fluctuations in the environment to relay information to the nervous system. Often times, receptors will sense multiple stimuli, such as gustatory receptors contributing to olfaction, light and temperature sensation (Montell, 2021). While mosquitoes are relatives of the well-studied model Dipteran, Drosophila melanogaster, their divergence to hematophagous behavior distinguishes them with a unique set of chemosensory receptors, including the critically important olfactory receptors.

            There are 3,559 species of mosquitoes characterized today (Harbach, 2018), few of which have had their olfactory systems examined comprehensively and at the molecular level. This is because few mosquito species are actually anthropophilic (Reeves et al., 2018) and many don’t feed on blood at all (Rattanarithikul et al., 2010), narrowing down the candidate genes responsible for human host seeking behavior. Olfactory perception starts at the peripheral organs in mosquitoes, which will be the areas of localized olfactory receptor expression and include the antennae, maxillary palps and proboscis in Anopheles (Saveer et al., 2018), Aedes (Lombardo et al., 2017) and Culex (Leal et al., 2013). Mosquito antennae are morphologically similar across species with antennae being bushier in males and more slender in females, although there are exceptions as in the Sabethes mosquitoes, which lack sexual dimorphism in the antennae and palps. Antennae can also detect sound (Su et al., 2018) so it is likely that their morphology has been driven by the evolutionary demands of olfaction and hearing. Maxillary palps show more morphological diversity between species and sex, with Anopheles males having long club-shaped maxillary palps and females having cylindrical palps that are slightly shorter, for example. Maxillary palps carry both chemosensory sensilla and mechanosensory bristles so that perception can be fine-tuned based on the array of molecularly differentiated sensilla on the palps (McIver and Hudson, 1972). The proboscis is more morphologically complex, made up of a pair of maxillae with teeth-like structures, a pair of mandibles, a needle-like labrum and a hypopharynx, all encased in a labium and ending in a labellum (Choo et al., 2015). Males do not blood feed and do not need to pierce the skin of a host with their proboscis, which is reflected in the transcriptomic profile of their proboscis and salivary glands. Interestingly, when C. quinquefasciatus males were introduced blood in their diet, they had significantly reduced lifespans, demonstrating the divergent evolutionary paths taken by different sexes (Nikbakhtzadeh, Buss and Leal, 2016).

The emergence and spread of Ae. albopictus

            Ae. albopictus, the Asian tiger mosquito is believed to have colonized the United States in the mid-1980s (Sprenger and Wuithiranyagool, 1986). Genetic examination of the mitochondrial NADH dehydrogenase subunit 5 (ND5) sequence indicate that Ae. albopictus originated in the Unites States from a small population of founder females and that genetic drift had insufficient time to reduce variation at nuclear loci (Birungi and Munstermann, 2002). Ae. albopictus has proven to be a robust invasive species that demonstrates ecological plasticity, being able to survive over winter and in cooler climates, as its eggs undergo a unique diapause (Armbruster, 2016). Their dissemination throughout the United States appears to be due to urbanization and climatic variables as they are not known to have a large flight range (Guerra et al., 2014). They are expected to increase their niche expansion into new climatically suitable urban areas beyond 2030 and especially 2050, which needs to be addressed if we are to combat the deadly vector-borne diseases that they transmit to humans (Kraemer et al., 2019). Ae. albopictus is known as a vector of 7 different alphaviruses, 8 bunyaviruses and 3 flaviviruses (Gratz, 2004) as well as the parasite responsible for Dirofilarial infection (Genchi et al., 2009).

            Ae. albopictus undergoes a semi-aquatic lifecycle with the larval and pupal stages taking place in stagnant pools of water. Laboratory studies show that adult females of Ae. albopictus reared at 25°C with a photoperiod of 16 hr 90% of their eggs hatched within 10 days while eggs from females reared at 21°C with a photoperiod of 10 hr only had a survival of 5% (Mori, Oda and Wada, 1981). A test to determine the nutritional requirements for optimal larval development determined that 5 mg of TetraMin (20.34 ± 0.03 kJ/g) produced the most robust adults (Müller et al., 2013).

            As Ae. albopictus is steadily increasing its geographic spread across the world and because it transmits multiple deadly human pathogens, many genetic methods have been examined as potential tools for vector control. Initial attempts at micro-injection of embryos were successful at producing germline mutations in Ae. albopictus, inserting a piggyBac-based transgene carrying a 3xP3-ECFP marker and an attP site (Labbé, Nimmo and Alphey, 2010). Transformation efficiencies were 2-3% and established five independent transgenic lines that facilitate secondary downstream transgenesis. A conditional female-specific late-acting flightless phenotype was later induced by using the Ae. albopictus Actin-4 gene to drive a dominant lethal gene in the indirect flight muscles (Labbé et al., 2012). Inducible dominant lethal genes can be used as genetic tools to create effectively sterile males and reduce populations in the field. Genetic tools for use in arthropods continue to expand and they must be thoroughly tested in the lab before they can be utilized in the field.

The development of molecular tools

            The biological sciences have been greatly impacted by technological advances in the past 25 years. Perhaps the most rapid advancement has been seen in DNA sequencing and bioinformatics, which has led to a deep understanding of the biology and genetics of non-model organisms relevant to public health, agriculture, and socioeconomics. Currently, complete genome annotations of over 150 insects have been made available on public databases with the majority coming from the Diptera and Hymenoptera (Li et al., 2019). This includes extensive population data to account for field diversity, made in efforts to control major epidemics like malaria (Miles et al., 2017). These advancements have made it possible to obtain sequencing and “omics” data that allows for the design of targeted gene engineering of organisms, facilitating the further exploration of gene function, genetic networks, and interactions between vectors and the pathogens they transmit.

            Genetic manipulation of arthropods of medical and economic importance is a persistent challenge within the entomology field. Vector genome modification techniques were standardized in the 1990s, with the development of expression vectors including baculovirus (Maeda et al., 1985), SINV (Higgs et al., 1995), plasmid (Cornel et al., 1997) and transposon-mediated integration (P-element (Miller et al., 1987), Hermes (Jasinskiene et al., 1998), Minos (Catteruccia et al., 2000), and piggyBac (Kokoza et al., 2001). These expression vectors were used to express foreign DNA in mosquitoes to study their biology, biocontrol, and to produce exogenous gene products. The DNA must be delivered to germ cells to create heritable genetic changes that will be transmitted through generations. Traditional DNA delivery techniques have relied on direct microinjection of early-stage embryos with gene-vectors that produce a random (piggyBac, Hermes, Minos, Mos1) or site-specific (phiC31, TALEN, zinc-finger nucleases) insertions into the genome. Major systems used for insect transgenesis include the piggyBac transposon (Handler and Ii, 1999; Grossman et al., 2001; Kokoza et al., 2001) and φC31 (Nimmo et al., 2006; Labbé, Nimmo and Alphey, 2010), which have been successful for a range of species, and for which reagents, such as insect lines that constitutively express PB transposase, φC31 recombinase, or contain AttP/AttB docking sites, are widely available. Other less technical methods like biolistics (Kravariti et al., 2001; Lule-Chávez et al., 2021) or electroporation (Thomas, 2003) that were originally developed for the manipulation of somatic tissues are now showing promise for germline transformation.

            The beginnings of CRISPR/Cas’s story with molecular biologists began in the late 1980s at the University of Alicante, Spain as a thesis project focused on the extremely halophilic archaea Haloferax mediterranei R-4 (Rodriguez-Valera, Juez and Kushner, 1983). When a scoop from a different research group shifted the team’s objectives, they began to explore unstudied regions of the genome of H. mediterranei that seemed to be subjected to some sort of salt-associated DNA modification (Juez et al., 1990). They discovered 30 base pair repeat regions flanked at one end by a highly degenerate copy (Mojica, Juez and Rodriguez‐Valera, 1993). It was eventually discovered to be a form of adaptive immunity in prokaryotes and archaea that defends against invading genetic elements. The repeat regions were initially called short regularly spaced repeats until 2012, when they were given their current appellation (Mojica and Garrett, 2013). After the initial discovery, many groups began to explore the role of the elusive repeat sequences and their associated genes, so it is difficult to give credit to any one team (Lander, 2016). Regardless, the utility of the CRISPR/Cas system as a molecular tool to target and introduce specific double-stranded mutations almost anywhere in the genome proved to be a monumental development in molecular biology.

            Cas proteins are contained in operons adjacent to the CRISPR repeats and contain different domains characteristic of nucleases, helicases, polymerases and various RNA-binding proteins (Jansen et al., 2002). The mechanisms of RNAi were being discovered prior to CRISPR/Cas and indeed there are certain specific, but evolutionary disparate, parallels between the two systems (Carthew and Sontheimer, 2009). The CRISPR/Cas system works through three phases: adaptation, expression and interference. During adaptation, invading genetic elements are incorporated upstream of the Cas and leader gene sequences and in between repeat sequences. During expression, they are transcribed into a pre-crRNA which includes many spacer and repeat sequences. This is further processed into the crRNA which acts as the template that targets complementary sequences and guides a Cas protein to degrade the targeted regions during the interference stage. Throughout the process, there are many different Cas proteins that play many different roles, some of which are still being discovered today (Thompson, Sobol and Prakash, 2021).

Objectives of this study:

            The major goals of this study were to leverage the most recent gene function interrogation methodologies in modulating the expression of functionally important arthropod genes. The overarching hypothesis was that developing such techniques would lead to a better understanding of gene function in blood feeding arthropods investigated in this study. (a) RNAi technique was used to knock-down salivary nitrophorins in C. lectularius and to record behavioral phenotypes involved in blood feeding. (b) A knock-out of a gene encoding an olfactory receptor in Ae. albopictus was developed using CRISPR/Cas9 while simultaneously knocking-in the dsRED marker coding sequence. These experiments build on the available toolsets used for the genetic modulation of arthropods and expand our understanding of molecular biology.

References

Aak, A. et al. (2014) ‘Questing activity in bed bug populations: male and female responses to host signals’. Physiol Entomol, 39(3), pp. 199–207. doi: 10.1111/PHEN.12062.

Alderton, W., Cooper, C. and Knowles, R. (2001) ‘Nitric oxide synthases: structure, function and inhibition’. Biochem J, 357(Pt 3), pp. 593–615. doi: 10.1042/0264-6021:3570593.

Anderson, A. L. and Leffler, K. (2008) ‘Bedbug infestations in the news: a picture of an emerging public health problem in the United States’, Journal of Environmental Health, 70(9), pp. 24–53.

Armbruster, P. A. (2016) ‘Photoperiodic diapause and the establishment of Aedes albopictus (Diptera: Culicidae) in North America’, Journal of Medical Entomology. Entomological Society of America, 53(5), pp. 1013–1023. doi: 10.1093/jme/tjw037.

Benoit, J. et al. (2011) ‘Drinking a hot blood meal elicits a protective heat shock response in mosquitoes’, Proceedings of the National Academy of Sciences of the United States of America. Proc Natl Acad Sci U S A, 108(19), pp. 8026–8029. doi: 10.1073/PNAS.1105195108.

Birungi, J. and Munstermann, L. E. (2002) ‘Genetic Structure of Aedes albopictus (Diptera: Culicidae) populations based on mitochondrial ND5 sequences: evidence for an independent invasion into Brazil and United States’, Annals of the Entomological Society of America, 95(1), pp. 125–132.

Calvo, E. et al. (2006) ‘Function and evolution of a mosquito salivary protein family’, Journal of Biological Chemistry, 281(4), pp. 1935–1942. doi: 10.1074/jbc.M510359200.

Carthew, R. W. and Sontheimer, E. J. (2009) ‘Origins and mechanisms of miRNAs and siRNAs’, Cell. Cell, 136(4), pp. 642–655. doi: 10.1016/j.cell.2009.01.035.

Catteruccia, F. et al. (2000) ‘Toward anopheles transformation: minos element activity in anopheline cells and embryos’, Proceedings of the National Academy of Sciences. National Academy of Sciences, 97(5), pp. 2157–2162. doi: 10.1073/PNAS.040568397.

Choo, Y. et al. (2015) ‘Multitasking roles of mosquito labrum in oviposition and blood feeding’, Frontiers in physiology. Front Physiol, 6(OCT). doi: 10.3389/FPHYS.2015.00306.

Cohen, A. C. (1995) ‘Extra-oral digestion in predaceous terrestrial arthropoda’, Annual Review of Entomology. Annual Reviews, 40(1), pp. 85–103. doi: 10.1146/annurev.en.40.010195.000505.

Cornel, A. et al. (1997) ‘Transient expression of the Drosophila melanogaster cinnabar gene rescues eye color in the white eye (WE) strain of Aedes aegypti’, Insect biochemistry and molecular biology. Insect Biochem Mol Biol, 27(12), pp. 993–997. doi: 10.1016/S0965-1748(97)00084-2.

Davies, T. G. . E., Field, L. M. and Williamson, M. S. (2012) ‘The re-emergence of the bed bug as a nuisance pest: implications of resistance to the pyrethroid insecticides’, Medical and Veterinary Entomology, 26(3), pp. 241–254. doi: 10.1111/j.1365-2915.2011.01006.x.

Doggett, S. L., Geary, M. J. and Russell, R. C. (2004) ‘The resurgence of bed bugs in Australia: with notes on their ecology and control’, Environmental Health, 4(2), pp. 30–38.

Ellis, A. M. et al. (2011) ‘Parameterization and sensitivity analysis of a complex simulation model for mosquito population dynamics, dengue transmission, and their control’, The American Journal of Tropical Medicine and Hygiene. The American Society of Tropical Medicine and Hygiene, 85(2), p. 257. doi: 10.4269/AJTMH.2011.10-0516.

Foote, A. et al. (2015) ‘Convergent evolution of the genomes of marine mammals’, Nature genetics. Nat Genet, 47(3), pp. 272–275. doi: 10.1038/NG.3198.

Francischetti, I. M. B. et al. (2010) ‘Insight into the sialome of the bed bug, Cimex lectularius.’, Journal of proteome research. NIH Public Access, 9(8), pp. 3820–31. doi: 10.1021/pr1000169.

Freitas, L. and Nery, M. F. (2020) ‘Expansions and contractions in gene families of independently-evolved blood-feeding insects’, BMC Evolutionary Biology 2020 20:1. BioMed Central, 20(1), pp. 1–8. doi: 10.1186/S12862-020-01650-3.

Gantner, B., LaFond, K. and Bonini, M. (2020) ‘Nitric oxide in cellular adaptation and disease’, Redox biology. Redox Biol, 34. doi: 10.1016/J.REDOX.2020.101550.

Gao, D. et al. (2016) ‘Prevention and control of Zika as a mosquito-borne and sexually transmitted disease: A mathematical modeling analysis’, Scientific Reports. Nature Publishing Group, 6. doi: 10.1038/SREP28070.

Genchi, C. et al. (2009) ‘Climate and Dirofilaria infection in Europe’, Veterinary Parasitology. Vet Parasitol, 163(4), pp. 286–292. doi: 10.1016/j.vetpar.2009.03.026.

Goddard, J. and DeShazo, R. (2009) ‘Bed bugs (Cimex lectularius) and clinical consequences of their bites’, JAMA. American Medical Association, 301(13), p. 1358. doi: 10.1001/jama.2009.405.

Goddard, J. and De Shazo, R. (2012) ‘Psychological effects of bed bug attacks (Cimex lectularius L.)’, American Journal of Medicine. Elsevier, 125(1), pp. 101–103. doi: 10.1016/j.amjmed.2011.08.010.

Gratz, N. G. (2004) ‘Critical review of the vector status of Aedes albopictus’, Medical and Veterinary Entomology. Med Vet Entomol, 18(3), pp. 215–227. doi: 10.1111/j.0269-283X.2004.00513.x.

Griesmann, M. et al. (2018) ‘Phylogenomics reveals multiple losses of nitrogen-fixing root nodule symbiosis’, Science. Science, 361(6398). doi: 10.1126/SCIENCE.AAT1743.

Grossman, G. et al. (2001) ‘Germline transformation of the malaria vector, Anopheles gambiae, with the piggyBac transposable element’, Insect molecular biology. Insect Mol Biol, 10(6), pp. 597–604. doi: 10.1046/J.0962-1075.2001.00299.X.

Guerra, C. A. et al. (2014) ‘A global assembly of adult female mosquito mark-release-recapture data to inform the control of mosquito-borne pathogens’, Parasites and Vectors. BioMed Central Ltd., 7(1). doi: 10.1186/1756-3305-7-276.

Handler, A. M. and Ii, R. A. H. (1999) ‘Germline transformation of Drosophila melanogaster with the piggyBac transposon vector’, Insect Molecular Biology. John Wiley & Sons, Ltd, 8(4), pp. 449–457. doi: 10.1046/J.1365-2583.1999.00139.X.

Harbach, R. E. (2018) Culicipedia: Species-group, genus-group and family-group names in Culicidae (Diptera). Cabi, Wallingford, UK. 2018, ISBN 9781786399052. CAB International.

Hentley, W. T. et al. (2017) ‘Bed bug aggregation on dirty laundry: A mechanism for passive dispersal’, Scientific Reports. Nature Publishing Group, 7(1), pp. 1–5. doi: 10.1038/s41598-017-11850-5.

Higgs, S. et al. (1995) ‘Mosquito sensitivity to a scorpion neurotoxin expressed using an infectious Sindbis virus vector’, Insect molecular biology. Insect Mol Biol, 4(2), pp. 97–103. doi: 10.1111/J.1365-2583.1995.TB00013.X.

Hwang, S. W. et al. (2005) ‘Bed bug infestations in an urban environment’, Emerging Infectious Diseases. Centers for Disease Control and Prevention (CDC), 11(4), pp. 533–538. doi: 10.3201/eid1104.041126.

Jansen, R. et al. (2002) ‘Identification of genes that are associated with DNA repeats in prokaryotes’, Molecular Microbiology. Mol Microbiol, 43(6), pp. 1565–1575. doi: 10.1046/j.1365-2958.2002.02839.x.

Jasinskiene, N. et al. (1998) ‘Stable transformation of the yellow fever mosquito, Aedes aegypti, with the Hermes element from the housefly’, Proceedings of the National Academy of Sciences of the United States of America. National Academy of Sciences, 95(7), p. 3743. doi: 10.1073/PNAS.95.7.3743.

Juez, G. et al. (1990) ‘Evidence for salt-associated restriction pattern modifications in the archaeobacterium Haloferax mediterranei’, Journal of Bacteriology. J Bacteriol, 172(12), pp. 7278–7281. doi: 10.1128/jb.172.12.7278-7281.1990.

Kokoza, V. et al. (2001) ‘Efficient transformation of the yellow fever mosquito Aedes aegypti using the piggyBac transposable element vector pBac [3xP3-EGFP afm]’, Insect Biochemistry and Molecular Biology. Pergamon, 31(12), pp. 1137–1143. doi: 10.1016/S0965-1748(01)00120-5.

Kraemer, M. U. G. et al. (2019) ‘Past and future spread of the arbovirus vectors Aedes aegypti and Aedes albopictus’, Nature Microbiology. Nature Publishing Group, 4(5), pp. 854–863. doi: 10.1038/s41564-019-0376-y.

Kravariti, L. et al. (2001) ‘The biolistic method as a tool for testing the differential activity of putative silkmoth chorion gene promoters’, Insect biochemistry and molecular biology. Insect Biochem Mol Biol, 31(4–5), pp. 473–479. doi: 10.1016/S0965-1748(00)00152-1.

Krinsky, W. L. (2019) ‘True Bugs (Hemiptera)’, in Medical and Veterinary Entomology. Academic Press, pp. 107–127. doi: 10.1016/B978-0-12-814043-7.00008-X.

Labbé, G. M. C. et al. (2012) ‘Female-specific flightless (fsRIDL) phenotype for control of Aedes albopictus’, PLoS Neglected Tropical Diseases. Public Library of Science, 6(7), p. e1724. doi: 10.1371/journal.pntd.0001724.

Labbé, G. M. C., Nimmo, D. D. and Alphey, L. (2010) ‘Piggybac- and PhiC31-mediated genetic transformation of the Asian tiger mosquito, Aedes albopictus (Skuse)’, PLoS Neglected Tropical Diseases. Public Library of Science, 4(8), p. e788. doi: 10.1371/journal.pntd.0000788.

Lander, E. S. (2016) ‘The Heroes of CRISPR’, Cell. Cell Press, 164(1–2), pp. 18–28. doi: 10.1016/j.cell.2015.12.041.

Leal, W. et al. (2013) ‘Differential expression of olfactory genes in the southern house mosquito and insights into unique odorant receptor gene isoforms’, Proceedings of the National Academy of Sciences of the United States of America. Proc Natl Acad Sci U S A, 110(46), pp. 18704–18709. doi: 10.1073/PNAS.1316059110.

Li, F. et al. (2019) ‘Insect genomes: progress and challenges’, Insect molecular biology. Insect Mol Biol, 28(6), pp. 739–758. doi: 10.1111/IMB.12599.

Lombardo, F. et al. (2017) ‘Deciphering the olfactory repertoire of the tiger mosquito Aedes albopictus’, BMC Genomics. BioMed Central, 18(1), pp. 1–23. doi: 10.1186/S12864-017-4144-1.

Lule-Chávez, A. et al. (2021) ‘Genetic transformation of mosquitoes by microparticle bombardment’, Insect molecular biology. Insect Mol Biol, 30(1), pp. 30–41. doi: 10.1111/IMB.12670.

Maeda, S. et al. (1985) ‘Production of human alpha-interferon in silkworm using a baculovirus vector’, Nature. Nature, 315(6020), pp. 592–594. doi: 10.1038/315592A0.

McIver, S. and Hudson, A. (1972) ‘Sensilla on the antennae and palps of selected Wyeomyia mosquitoes’, Journal of Medical Entomology. Oxford Academic, 9(4), pp. 337–345. doi: 10.1093/JMEDENT/9.4.337.

Miles, A. et al. (2017) ‘Genetic diversity of the African malaria vector Anopheles gambiae’, Nature. Nature, 552(7683), pp. 96–100. doi: 10.1038/NATURE24995.

Miller, L. et al. (1987) ‘Stable integration and expression of a bacterial gene in the mosquito Anopheles gambiae’, Science (New York, N.Y.). Science, 237(4816), pp. 779–781. doi: 10.1126/SCIENCE.3039658.

Mojica, F. J. M. and Garrett, R. A. (2013) ‘Discovery and seminal developments in the CRISPR field’, in CRISPR-Cas Systems. Springer Berlin Heidelberg, pp. 1–31. doi: 10.1007/978-3-642-34657-6_1.

Mojica, F. J. M., Juez, G. and Rodriguez‐Valera, F. (1993) ‘Transcription at different salinities of Haloferax mediterranei sequences adjacent to partially modified PstI sites’, Molecular Microbiology. Mol Microbiol, 9(3), pp. 613–621. doi: 10.1111/j.1365-2958.1993.tb01721.x.

Mol, C. et al. (1995) ‘Structure and function of the multifunctional DNA-repair enzyme exonuclease III’, Nature. Nature, 374(6520), pp. 381–386. doi: 10.1038/374381A0.

Montell, C. (2021) ‘Drosophila sensory receptors—a set of molecular Swiss Army Knives’, Genetics. Oxford Academic, 217(1), pp. 1–34. doi: 10.1093/GENETICS/IYAA011.

Mori, A., Oda, T. and Wada, Y. (1981) ‘Studies on the egg diapause and overwintering of Aedes albopictus in Nagasaki.’, Tropical Medicine, 23(2), pp. 79–90.

Müller, R. et al. (2013) ‘Appropriate larval food quality and quantity for aedes albopictus (diptera: Culicidae)’, Journal of Medical Entomology. Oxford Academic, 50(3), pp. 668–673. doi: 10.1603/ME12094.

Nikbakhtzadeh, M., Buss, G. and Leal, W. (2016) ‘Toxic effect of blood feeding in male mosquitoes’, Frontiers in physiology. Front Physiol, 7(JAN). doi: 10.3389/FPHYS.2016.00004.

Nimmo, D. et al. (2006) ‘High efficiency site-specific genetic engineering of the mosquito genome’, Insect molecular biology. Insect Mol Biol, 15(2), pp. 129–136. doi: 10.1111/J.1365-2583.2006.00615.X.

Nyanjom, S. et al. (2018) ‘Expression levels of odorant receptor genes in the savanna Tsetse Fly, Glossina morsitans morsitans’, Journal of medical entomology. J Med Entomol, 55(4), pp. 855–861. doi: 10.1093/JME/TJY018.

Omori, N. (1941) ‘Comparative studies on the ecology and physiology of common and tropical bed bugs, with special reference to the reactions to temperature and moisture.’, Journal of the Medical Association of Formosa, 40(3), pp. 555–636.

Paim, R. et al. (2016) ‘Functional evaluation of Heat Shock Proteins 70 (HSP70/HSC70) on Rhodnius prolixus (Hemiptera, Reduviidae) physiological responses associated with feeding and starvation’, Insect biochemistry and molecular biology. Insect Biochem Mol Biol, 77, pp. 10–20. doi: 10.1016/J.IBMB.2016.07.011.

Partha, R. et al. (2019) ‘Robust method for detecting convergent shifts in evolutionary rates’, Molecular biology and evolution. Mol Biol Evol, 36(8), pp. 1817–1830. doi: 10.1093/MOLBEV/MSZ107.

Potter, M. F. (2004) ‘Bedbugs: understanding and control’, Pest Control Technology, 32(8), pp. 13–17.

Potter, M. F. (2006) ‘The perfect storm: an extension view on bed bugs’, American Entomologist, 52(2), pp. 102–104.

Potts, R. et al. (2021) ‘Molecular analysis of the blood meals and bacterial communities of bed bugs (Cimex lectularius L.) to assess interactions with alternative hosts’, Parasitology Research. Springer Science and Business Media Deutschland GmbH, 120(4), pp. 1209–1217. doi: 10.1007/s00436-021-07079-x.

Poulos, T. L. (1996) ‘The role of the proximal ligand in heme enzymes’, JBIC Journal of Biological Inorganic Chemistry 1996 1:4. Springer, 1(4), pp. 356–359. doi: 10.1007/S007750050064.

Rattanarithikul, R. et al. (2010) ‘Illustrated keys to the mosquitoes of Thailand V. Genera Orthopodomyia, Kimia, Malaya, Topomyia, Tripteroides, and Toxorhynchites’, The Southeast Asian journal of tropical medicine and public health, 41 Suppl 1, pp. 1–225.

Reeves, L. E. et al. (2018) ‘Identification of Uranotaenia sapphirina as a specialist of annelids broadens known mosquito host use patterns’, Communications Biology 2018 1:1. Nature Publishing Group, 1(1), pp. 1–8. doi: 10.1038/s42003-018-0096-5.

Ribeiro, J. M. C. et al. (1993) ‘Reversible binding of nitric oxide by a salivary heme protein from a bloodsucking insect’, Science, 260(5107), pp. 539–541. doi: 10.1126/SCIENCE.8386393.

Rodriguez-Valera, F., Juez, G. and Kushner, D. J. (1983) ‘Halobacterium mediterranei spec, nov., a New Carbohydrate-Utilizing Extreme Halophile’, Systematic and Applied Microbiology. Syst Appl Microbiol, 4(3), pp. 369–381. doi: 10.1016/S0723-2020(83)80021-6.

Rosenfeld, J. A. et al. (2016) ‘Genome assembly and geospatial phylogenomics of the bed bug Cimex lectularius’, Nature Communications. Nature Publishing Group, 7, p. 10164. doi: 10.1038/ncomms10164.

Sackton, T. B. and Clark, N. (2019) ‘Convergent evolution in the genomics era: new insights and directions’, Philosophical Transactions of the Royal Society B: Biological Sciences. The Royal Society, 374(1777). doi: 10.1098/RSTB.2019.0102.

Sailer, R. (1952) ‘The bedbug. An old bedfellow that’s still with us.’, Pest Control. Springer, 10(20), pp. 22–72. doi: 10.1007/s00436-012-2862-5.

Saveer, A. et al. (2018) ‘Characterization of chemosensory responses on the Labellum of the malaria vector mosquito, Anopheles coluzzii’, Scientific reports. Sci Rep, 8(1). doi: 10.1038/S41598-018-23987-Y.

Scarpino, S. V. and Althouse, B. M. (2019) ‘Uncovering the hidden cost of bed bugs’, Proceedings of the National Academy of Sciences, 116(15), pp. 7160–7162. doi: 10.1073/pnas.1902404116.

Siva-Jothy, M. T. et al. (2019) ‘Female bed bugs (Cimex lectularius L) anticipate the immunological consequences of traumatic insemination via feeding cues’, Proceedings of the National Academy of Sciences of the United States of America. National Academy of Sciences, 116(29), pp. 14682–14687. doi: 10.1073/pnas.1904539116.

Sprenger, D. and Wuithiranyagool, T. (1986) ‘The discovery and distribution of Aedes albopictus in Harris County, Texas’, Journal of American Mosquito Control Association, 2(2), pp. 217–219.

Su, M. et al. (2018) ‘Sex and species specific hearing mechanisms in mosquito flagellar ears’, Nature communications. Nat Commun, 9(1). doi: 10.1038/S41467-018-06388-7.

Thomas, J. (2003) ‘Electroporation, an alternative to biolistics for transfection of Bombyx mori embryos and larval tissues’, Journal of insect science (Online). J Insect Sci, 3(1). doi: 10.1093/JIS/3.1.17.

Thompson, M. K., Sobol, R. W. and Prakash, A. (2021) ‘Exploiting DNA endonucleases to advance mechanisms of DNA repair’, Biology. Multidisciplinary Digital Publishing Institute  (MDPI), 10(6), p. 530. doi: 10.3390/biology10060530.

Titschack, E. (1930) ‘Untersuchungen über das wachstum, den nahrungsverbrauch und die eierzeugung. III. Cimex lectularius L.’, Zeitschrift für Morphologie und Ökologie der Tiere. Springer-Verlag, 17(3), p. 551. doi: 10.1007/BF00419296.

Tsujishita, Y. et al. (2001) ‘Specificity determinants in phosphoinositide dephosphorylation: crystal structure of an archetypal inositol polyphosphate 5-phosphatase’, Cell. Cell, 105(3), pp. 379–389. doi: 10.1016/S0092-8674(01)00326-9.

Usinger, R. L. (1966) Monograph of Cimicidae (Hemiptera - Heteroptera). College Park: Entomological Society of America.

Valenzuela, J. G. and Ribeiro, J. M. (1998) ‘Purification and cloning of the salivary nitrophorin from the hemipteran Cimex lectularius.’, The Journal of experimental biology, 201(Pt 18), pp. 2659–64.

Walker, A. et al. (2016) ‘Venoms of heteropteran insects: A treasure trove of diverse pharmacological toolkits.’, Toxins. MDPI AG, 8(2), pp. 43–43. doi: 10.3390/TOXINS8020043.

Weichsel, A. et al. (2005a) ‘Heme-assisted S-nitrosation of a proximal thiolate in a nitric oxide transport protein’, Proceedings of the National Academy of Sciences, 102(3), pp. 594–599. doi: 10.1073/pnas.0406549102.

WHO (2020) Fact sheet: vector-borne diseases.

Yoon, K. A. et al. (2021) ‘Comparative analyses of the venom components in the salivary gland transcriptomes and saliva proteomes of some heteropteran insects’, Insect Science. John Wiley & Sons, Ltd, 0, pp. 1–19. doi: 10.1111/1744-7917.12955.

PART I 

CHARACTERIZATION OF THE COMPLETE SALIVARY TRANSCRIPTOME IN CIMEX LECTULARIUS USING NEXT-GENERATION SEQUENCING

Background

            The interface between the arthropod ectoparasite and the vertebrate host represents a dynamic evolutionary struggle that plays out with a diverse set of molecular effectors on both sides. The vertebrate must maintain haemostasis to prevent blood loss and ward off potential pathogens, while the parasite must obtain blood to complete its lifecycle and pass on its genes to the next generation. Bed bugs (Cimex lectularius L.) are an important parasite of humans that have persisted through written history and remain a nuisance in many urban areas throughout the world. They cause an acute immune reaction known as cimicosis in humans but are not regarded as an important vector of disease, although there is evidence that they can harbor human pathogens (Peta and Pietri, 2021). As C. lectularius is a successful parasite of humans, a closer examination of its salivary gland transcriptome will offer insight into pharmacologically relevant molecular effectors in humans. Additionally, although it is not considered a human disease vector, it is host to many viruses and bacteria that could potentially emerge as zoonotic human pathogens (Zhang et al., 2020). This chapter will focus on defining the salivary repertoire of C. lectularius, and how these components effect its role as an ectoparasite in humans.

            Early morphological and physiological studies of hemipteran salivary glands date back to the early 1940’s (Baptist, 1941), which were followed by preliminary biochemical studies (Berridge and Patel, 1968), but it wasn’t until the turn of the 21st century that attempts were made to characterize complete molecular profiles of insect salivary transcripts and proteins, as was done in the malaria parasite vector, Anopheles gambiae, whose cDNA profile was partially cataloged in 2002 (Francischetti et al., 2002). The first preliminary sialome in C. lectularius was published in 2010 (Francischetti et al., 2010b) and predicted the presence of nearly 100 different proteins in bed bug saliva. Through elucidation of the immunomodulatory effectors inducing vertebrate immunity, we can gain insight into allergic reactions and develop methods to counteract them. Additionally, salivary proteins and transcripts can play roles in immunity, homeostatic regulation or both (Chen et al., 2020). However, hematophagy has evolved independently in at least five different orders of insects so it is difficult to draw conclusions based on comparative genomics (Grimaldi and Engel, 2005). Because of this, it is necessary to make functional classifications of proteins based on the unique sequence domains within an individual species by first sequencing the gene or genome and then defining the transcriptional profile of individual tissue types.

            The first salivary transcriptome in C. lectularius relied on laborious expressed sequence tag (EST) library prep and Sanger sequencing, which can lack sequencing depth and completeness (Francischetti et al., 2010b). They isolated mRNA from bed bug salivary glands, ligated the corresponding cDNA into λ TriplEx2 vectors and infected XL1- Blue Escherichia coli cells to make an EST library, which was ultimately sequenced on an ABI 3730xL DNA Analyzer. The 3730xL is a fully automated system that uses Sanger sequencing technology to produce a maximum of 1 million bases per day, which is 5 orders of magnitude less than is possible today with the Illumina NextSeq 2000 at about 100 billion bases per day. The resulting alignment produced 837 unique and 123 related EST clusters representing cDNAs within the salivary gland of C. lectularius. Of these, 47% were housekeeping genes, 44% were unknown, 7% were secreted and 2% were microbial, viral, or transposable elements. Of the 698 putatively secreted ESTs, 42% belonged to odorant-binding proteins and 37% were for the unique Cimex nitrophorin. The remaining secreted genes included Serpins, antigen-5 family members, lysozymes, a Cimex-type apyrase, serine proteases and esterases.

            Most of the proteins found in the saliva by Francischetti et al, 2010a, could be functionally categorized as enzymes. The salivary inositol polyphosphate phosphatase (IPPase) family IPPases includes Cimex nitrophorin and are normally ubiquitous intracellular enzymes involved with inositol phosphate catabolism and cell signaling (Harris et al., 2008). Similar secreted salivary IPPases have been identified in Rhodnius prolixus (Ribeiro et al., 2004), Triatoma brasiliensis (Santos et al., 2007) and T. infestans (Assumpção et al., 2008) but are notably lacking in other pervasive hematophagous species like mosquitoes and ticks. The salivary Cimex nitrophorin carries a heme ligand that delivers NO to the host, which acts as a vasodilator and keeps blood flowing to the bite site (Valenzuela, Walker and Ribeiro, 1995). A total of 8 nitrophorins classified by sequence alignment were identified in the saliva of C. lectularius, which were phylogenetically divided into 2 subclades (Francischetti et al., 2010b). A polyacrylamide gel electrophoresis (PAGE)/mass spectrometry (MS)/MS band indicated the presence of an abundant protein of 32 kDa, which is consistent with the weight of salivary Cimex nitrophorins. There were 15 ESTs that had a 95% or higher alignment to a previously described apyrase (Valenzuela et al., 1998) that were additionally present at the predicted 39 kDa weight on the PAGE/MS/MS. These ESTs were all predicted to be from a single copy gene. The salivary apyrase acts as an inhibitor of platelet aggregation, facilitating blood flow to the bite site (Valenzuela, Chuffe and Ribeiro, 1996). Two genes that had a 60% alignment to each other represented 8 ESTs and belonged to the Diadenosine and diphosphoinositol polyphosphate phosphohydrolase family of enzymes. The diadenosine phosphatases are believed to be responsible for signal transduction and may play a role in the hydrolysis of proinflammatory diadenosine nucleotides Ap4A and Ap5A, which are released by platelets and neutrophils (Stavrou, 2003). Forty of the ESTs found in the Cimex salivary library align closely to the carboxy terminus domain of insect proteins classified as esterases or cholinesterases (Francischetti et al., 2010b). This enzyme family has been identified in other hematophagous arthropods but its role in blood feeding is unclear, however acetylcholinesterase genes have been implicated with insecticide resistance (Atyame et al., 2019). Within the serine protease family of enzymes, trypsin or chymotrypsin-like enzymes are commonly found in hematophagous arthropod salivary glands (Ribeiro and Arcà, 2009) and they were represented by 18 ESTs in the Francischetti et al, 2010a, study.

Materials and methods

1.1.1             Maintenance and collection of colony arthropods

Cimex lectularius Harlan strain were maintained in 4 oz glass mason jars with an 8 cm mouth, 6.5 cm base and 6.5 cm height sealed by a metal canning lid. The lids rubber-gasketed insert was removed and replaced with nylon netting weaved to 0.5 mm by 0.5 mm squares. All jars contained two cardboard, single layer cardstocks, folded on itself approximately every 0.5 cm. All cardboard was scraped with a sharp edge to increase surface roughness and promote oviposition by females. Jars, lids and cardboard was autoclaved before the first use as they would remain with that particular colony throughout the course of their lives. The jars were assembled so that the folded cardboard extended to the top of the 6.5 cm jar and contacted the netting. This ensured that the bed bugs could feed from membrane feeders placed on the top of the upright jars. Membrane feeds were done using feeders connected to a 37°C water bath by Clippard 0.25 OD, 0.125 ID polyurethane tubing (URH1-0804) and sealed with a 4 cm by 4 cm square piece of Bevis Parafilm (PM-999) stretched over the surface of the feeder in contact with the netting. A P1000 pipette was used to load the membrane feeders from the top using 0.5 ml of rabbit blood containing 3.4% sodium citrate (HemoStat Laboratories; #RBC025). Bed bugs were allowed to feed for approximately 30 minutes to 1 hour until the majority have fed to repletion. For long term storage of C. lectularius, jars were stored at 10°C for up to 400 days (Omori, 1941). Bed bugs that were

Figure 1.1           Creating the colony jar for C lectularius. (a) The netting must not be large enough by squared area to allow for any 1st stage nymphs to escape. The netting must be free of toxic additives and not affixed to the jar with super glue as this will kill the bed bug. (b) The netting is glued to the canning lid with hot glue. The insert to the canning lid is discarded and a ring of glue is applied around the circumference of the lid from the inside. The outside is covered with hot glue at the interface with the netting. Continued freeze-thaw cycles with the jar will crack the hot glue and require re-application. (c) The finished jar with folded cardboard insert. The cardboard or paper stock should be rubbed with an abrasive to promote oviposition by female bed bugs.

being used actively for experiments were stored at 30°C in an EchoTherm in40 chilling incubator to minimize the time spent in each nymphal stage. At 30°C eggs hatch in, on average, 4.4 days, 1st instar nymphs transition to 2nd instars in 4.4 days, 2nd to 3rd in 2.8 days, 3rd to 4th in 2.4 days, 4th to 5th in 3.2 days and 5th instar to adult in 4 days for a total time to adulthood of 24.2 days (Omori, 1941). Adults were harvested from the jars under a fume hood and direct light on top of a large serving tray. The edge of the serving tray was lined with a thick layer of petroleum jelly to prevent the escape of any bed bugs. Adult collections were done by tilting the jars into a collection container with the non-dominant hand while using a paint brush to move individuals crawling towards the mouth of jar into a transfer vessel with the dominant hand. The sex of C. lectularius individuals was distinguished by a narrower, more pointed abdomen, versus a broader, more circular abdomen in males versus females, respectively.

Figure 1.2           Differentiating between male and female individuals of C. lectularius. The individual on the left is a female. The dotted yellow crescent outlines the ectospermalege, which is the external organ penetrated by the make during traumatic insemination. The posterior paratergite segment is more rounded than the male as outlined by the dotted black crescent line. The male is displayed on the right and the terminal abdominal paratergite is more pointed giving them a more oblong appearance when viewed dorsally.

1.1.2             Dissection of salivary tissue

            Microscope work was done using a Zeiss Discovery.V12 Stereoscope with a PlanApo S 0.63x FWD 81 mm lens and PI 16x/16 eyepiece (#444054-9000). The microscope photos were taken with a Zeiss Axiocam 208 color connected to Samsung monitor through an HDMI connection. All bed bugs were placed in a petri dish on ice to anesthetize them prior to any dissections to prevent escape. A custom dissection stage was made using a Globe Scientific Inc 25 mm by 75 mm by 1 mm glass microscope slide (#1304) and hot glue to facilitate a liquid working area that could be pinned. Using 1 cm metal pins, individual C. lectularius samples were affixed to the slide through the dorsal side of the 5th paratergite segment and the base of the clypeus. The working area was filled with phosphate buffered saline (PBS) so that the salivary glands remained

Figure 1.3           The dissection station used for salivary gland extraction. (a) The Zeiss Discovery.V12 Stereoscope with a PlanApo S 0.63x FWD 81 mm lens and PI 16x/16 eyepiece was used for all dissections. The stage is affixed to the base with blue tape to prevent movement of specimens and facilitate hand placement for dissection. (b) The custom glass slide created to allow for liquid dissections and salivary gland removal. The copious application of hot glue prevents the penetration of liquid, which will undermine the effectiveness of the glues attachment to the glass slide. (c) The dissection of the salivary gland from C. lectularius showing the gland outlined in a white dotted oval. Note the pronotums lateral fracture made with the scalpel to facilitate locating the salivary glands.

 in solution and could be easily removed. A carbon steel, feather double edge blade (Electron Microscopy Sciences; #72002-01) in blade clamps was used to make an incision across the entire dorsal surface between the pronotum and the mesonotum-scutellum. The pronotum was lifted until the two salivary glands could be removed using dissecting jewelers microforceps (Fisherbrand; #08-953E). The salivary glands appeared bright orange or pink and were placed into a 1.5 ml microcentrifuge tube containing 200 ml of RNA later (Sigma Life Sciences; Cat #R0901) and were incubated at -80°C until RNA extraction. The remaining portions of the carcass were unpinned, removed from the PBS, placed in 200 ml of RNAlater and stored at -80°C until RNA extraction.

Figure 1.4           A representation of the dorsal view of the dorsal anatomy of C. lectularius. The cartoon on the left displays the bed bug with the right hemelytral removed to display the underlying anatomy. A corresponding image of a male specimen is displayed on the right. The image was taken with Zeiss Discovery.V12 Stereoscope under 32x magnification and the right hemelytral was dissected to expose the underlying anatomy.

Figure 1.5           The salivary gland of Cimex lectularius. A) The location of the salivary glands can be vaguely determined by their pink/orange color near the junction of the head and pronotum in a normally pigmented individual.  The glands are circled in the inset of the more intensely backlit image. B) Light microscope image showing the orange-pink color of the heme protein, nitrophorin. B) Fluorescence microscopy showing the musculature stained with Phalloidin (green) and nuclei stained with DAPI (blue) of the same salivary gland.

1.1.3             Total RNA extraction

            Total RNA was extracted from the salivary glands and carcasses of 80 adult females using the Purelink mini kit (Thermo Fisher; Cat #12183018A). Tissues were removed from the -80°C freezer and thawed on ice. They were centrifuged on a Sorvall Legend Micro17 Thermo Fisher centrifuge at 4°C and 16,000 x g for 1 min and the RNAlater supernatant was carefully removed from the tissue pellet on ice. The tissues were quickly resuspended in 3 µl of 2-mecaptoethanol and 0.3 ml of Lysis Buffer. The tissues were homogenized using an RNase-free pestle using an up and down twisting motion directed towards the bottom of the tube until the tissues were completely disrupted. The homogenate was centrifuged for 2 min at 12,000 x g and the supernatant was transferred to a clean RNase-free 1.5 ml microcentrifuge tube. The lysate was then mixed with 400 µl of 100% ethanol and vortexed thoroughly. This mixture was pipetted into a spin column inserted into a collection tube and centrifuged at 12,000 x g at room temperature for 15 sec. The flow through was removed and 700 µl of Wash Buffer 1 was added to the spin column and the tube was spun at 12,000 x g for 15 sec at room temperature. The collection tube was discarded with the flow through and replaced. 500 µl of Wash Buffer 2 was added to the column and centrifuged at 12,000 x g for 15 sec at room temperature and the flow through was discarded. The wash steps were repeated. A final drying step was carried out by centrifuging the column with collection tube at 12,000 x g for 1 min at room temperature. The collection tube was discarded, and the spin column was placed into a clean RNase-free 1.5 ml microcentrifuge tube. Elution was done by adding 30 µl of RNase-free water directly to the column and incubating at room temperature for 2 min. The spin column was then spun at 12,000 x g for 2 min at room temperature. The purified RNA was quantified using the Thermo Scientific Nanodrop One-C Microvolume UV Spectrophotometer and Quality of RNA was assessed using a Bioanalyzer 2100 (Agilent).

1.1.4             Illumina sequencing

            Total RNA was sent to the Hudson Alpha Genomic Services Laboratory (Huntsville, Alabama) for library preparation and directional RNA sequencing targeting at least 50 million paired-end reads of 50 base pairs. Poly(A) mRNA was isolated using the NEBNext Poly(A) mRNA Magnetic Isolation Module and the NEBNext Ultra Directional RNA Library Prep Kit (NEB) was used for library prep. First-strand synthesis of RNA was performed with actinomycin D included to inhibit DNA-dependent DNA synthesis. During second strand synthesis, dUTP was substituted for dTTP to ensure that only the first strand was synthesized, conferring directionality and strandedness. The libraries were sequenced using the Illumina HiSeq 4000 platform.

1.1.5             Differential transcriptome analysis

            After obtaining the raw reads in fastq format, quality was assessed using FastQC version 0.11.4 (Andrews et al., 2010). Reads were trimmed using Trimmomatic version 0.36 (Bolger, Lohse and Usadel, 2014) with a quality threshold of 30 and a minimum read length of 50 to remove low quality base pairs. Only reads in which both pairs survived trimming were used. The reference genome and annotation files for C. lectularius were downloaded from NCBI in fna and gff formats, respectively. HISAT2 version 2.1.0 (Kim, Langmead and Salzberg, 2015) was used to generate a reference index of the C. lectularius genome (Jeffrey A Rosenfeld et al., 2016). Trimmed reads were then aligned to the reference index with default parameters to generate a sam file for each dataset (salivary and carcass). These sam files were then converted to bam format and sorted using Samtools version 1.8 (Li et al., 2009) with the flags view and sort, respectively. Sorted bam files were then assembled using StringTie version 1.3.0 (Pertea et al., 2016) with default parameters. The assembled gtf files were then merged using StringTie with the merge flag to generate a single assembly gtf file. A final set of StringTie assemblies involved assembling the sorted bam files with the merged assembly gtf as a reference. These assemblies were then converted to an EdgeR compatible format using a Python script supplied by StringTie. These assemblies were then imported to R-Studio and analyzed using the R package EdgeR version 3.20.9 (Robinson, McCarthy and Smyth, 2010). Relative expression was calculated using the estimateDisp function alongside the exactTest function. These data were formatted for easier viewing and plotting using standard R commands. An MA (log ratio-mean average) plot was generated using the R package ggplot2. Genes that were found to belong to the groups previously noted as important were colored as indicated in the legend. Gene function was found primarily using the reference annotation file, and the remaining genes were found by using the BLAST algorithm through NCBI to match genes of interest in closely related genes to those in C. lectularius. A more robust annotation was obtained using the program Blast2GO, which also generated Gene Ontology terms and statistics.

Results

1.1.6             Transcriptome analysis and assembly

The dataset for the bed bug carcasses contained 82,656,328 paired reads before trimming, of which 58,961,837 (71.33%) survived trimming and were used in further analysis. The dataset for the bed bug salivary glands contained 89,605,908 paired reads before trimming, of which 64,721,981 (72.23%) survived trimming and were used in further analysis. The carcass and salivary gland datasets had overall alignment rates of 79.97% and 83.15%, respectively. Detailed alignment statistics can be found in supplemental information. 19,269 unique genes were identified, of these 10,587 (54.94%) were differentially expressed with an alpha of 0.01. To reduce the number of genes to analyze, only those that were upregulated or downregulated at least 2-fold were considered differentially expressed for the remaining analysis. This left 7,930 genes upregulated in the salivary glands compared to the carcass, and 2,657 downregulated in the salivary glands compared to the carcass (Fig 2; S1 Table).

Figure 1.6         Plot of the log ratio versus mean average (MA) of salivary RNA levels vs. whole body minus salivary gland tissue. Each spot represents the fold change between tissues and mean expression values of one gene. Gene families of interest are color-coded. ACHE, Acetylcholinesterase; IPP5P, Inositol polyphosphate 5-phosphatases; Serpin, serine protease inhibitor; OBP, Odorant binding protein family. Light gray points are not considered differentially expressed, with a p-value greater than the set alpha value of 0.01, while darker gray points are considered differentially expressed, with a p-value less than the set alpha values of 0.01. Dark points lying the region above the non-differentially expressed center (light) are enriched in the salivary tissue versus the body without salivary glands. The box in the top right shows the area of highest interest for downstream analyses and discussion.

1.1.7             Characterization of major salivary proteins in C. lectularius

            Table 1 gives a summary of important salivary genes with the highest enrichment in the salivary gland of C. lectularius according to their log2-fold change. The list of gene includes two apyrases, five acetylcholinesterases, twelve lysozymes, seven odorant binding proteins, four serpins, two nudix hydrolases and eight IPP5Ps. Of the highly expanded lysozyme family, five are dominant in the bed bug sialome and branch off from Lepidopteran lysozymes (Fig 3). Two lysozymes with conserved glutamic and aspartic acid residues group with immune-related lysozymes in Harmonia and are believed to have muramidase activity. The normally constitutively expressed intracellular IPP5P genes have expanded and display disparate functionalities in the salivary gland of C. lectularius (Fig 4). These include the well described salivary nitrophorins and four closely related IPP5P-like genes, three of which have a clear signal peptide.

Table 1.1 Summary of the major salivary genes in C. lectularius

Gene name - GeneID - Annotation - Signal - peptide - Log2FC

ClecNitro1 - 106662976 - 72 kDa inositol polyphosphate 5-phosphatase-like - yes - 16.40

 ClecNitro2 - 106662977 - 72 kDa inositol polyphosphate 5-phosphatase-like - yes - 15.79

ClecNitro-like1 - 106663254 - inositol polyphosphate 5-phosphatase OCRL-1-like - yes - 16.28

ClecNitro-like2 - 106662975 - 72 kDa inositol polyphosphate 5-phosphatase-like - yes - 13.88

ClecNitro-like3 - 106662978 - inositol polyphosphate 5-phosphatase K-like - yes - 13.46

ClecNitro-like4 - 106662979 - inositol polyphosphate 5-phosphatase K-like - yes - 13.94

ClecNudix - 106666860 - bis(5'-nucleosyl)-tetraphosphatase [asymmetrical]-like - yes - 20.08

ClecEsterase1 - 106669436 - acetylcholinesterase-like - no - 16.68

ClecEsterase2 - 106673686 - acetylcholinesterase-like - no - 17.02

ClecEsterase3 - 106669437 - acetylcholinesterase-like - no - 16.81

ClecApyrase1 - 106669828 - apyrase - no - 12.63

ClecLysozyme1 - 106663584 - lysozyme-like - yes - 15.76

ClecLysozyme2 - 106666694 - lysozyme-like - yes - 16.83

ClecLysozyme3 - 106663588 - lysozyme c-1-like -yes - 12.90

ClecLysozyme4 - 106669094 - lysozyme c-1-like - yes - 16.30

ClecLysozyme5 - 106667626 - lysozyme c-1-like - no - 13.30

Figure 1.7 Expansion in the non-muramidase group of lysozyme genes in Cimex lectularius including five proteins highly expressed in the salivary glands[1].

Figure 1.8 The Cimex lectularius genome contains two functioning Inositol polyphosphate 5-phosphatases (IPP5P) genes, like other metazoans, along with six other IPP5P-related genes including two salivary nitrophorins and four other genes that are highly expressed in the salivary tissue and group phylogenetically with salivary nitrophorin[2].

1.1.8 Structural models of the major salivary proteins

            Using the ClecNitro1 crystal structure, 1y21.1.A, previously determined (Weichsel et al., 2005a) as the template query, models of the 5 other nitro and nitro-like proteins were made to assess potential functionalities (Table 2). Manually looking at the amino acids comprising the heme-binding pocket of these 5 nitrophorins can determine whether they are suitable for heme conjugation. Notably, only ClecNitro2 shares the critical Cys-60 necessary for heme conjugation and NO binding. Based purely on the Swiss-Modeling, none of the 5 nitrophorin models contain the heme protoporphyrin ring as the binding site is not conserved. This indicates that there is deviation from 100% identity between the query and template at the binding site and may not be an accurate characterization of associated ligands.

Table 1.10 Modeled alignment of the CleNitro proteins

Gene name - Template - Sequence identity - GMQE - QMEAN

ClecNitro1 - 1y21.1.A - 100.00% - 0.88 - 0.62

ClecNitro2 - 1y21.1.A - 85.41% - 0.86 - 0.37

ClecNitro-like1 - 1y21.1.A - 31.52% - 0.61 - (-3.26)

ClecNitro-like2 - 1y21.1.A - 29.09% - 0.59 - (-2.58)

ClecNitro-like3 - 1y21.1.A - 31.52% - 0.61 - 3.26

ClecNitro-like4 - 1y21.1.A - 31.16% - 0.61 - (-3.60)

Discussion

            These data corroborate and expand our current understanding of the C. lectularius sialome (Francischetti et al., 2010a) by using RNAseq with a recently released reference genome (Rosenfeld, et al., 2016). There are 391 bed bug salivary transcripts present at levels ~4-fold higher (log2 15 - log2 20) than their whole-body counterparts and include significant proteins previously identified in hematophagous arthropod saliva, including an apyrase, acetylcholinesterase and serpin. IPP5Ps and lysozymes make up most salivary proteins and were here characterized functionally based on phylogenetic analysis and conserved structural motifs. Because C. lectularius represents an ancient lineage that evolved hematophagous behavior independently within the Cimicomorpha, it is difficult to make assumptions about protein function based solely on homology. Selection pressure among salivary proteins is rapid and is higher within species utilizing disparate hosts than it is within species in different geographies (Talbot et al., 2017). The main advantage of a reference based RNAseq analysis is our confidence in sequence identity and quantity, especially with closely related paralogs. Without experimental evidence on individual proteins, we must infer function based on previously published empirical evidence and the biochemistry within the parasite/host blood feeding interface.

            Proteins in the IPP5P family dominate the salivary glands in both abundance and functionality. These salivary lipocalins are believed to have originated from gene duplication and neofunctionalization events (Wang et al., 2016) and are functionally unrelated to their constitutively expressed, intracellular counterparts. I identified six salivary enriched IPP5Ps, which are believed to function as phosphatases and nitrophorins. The Cys-60 of two Cimex salivary nitrophorins (LOC106662976 and LOC106662977) first conjugate with ferric heme in the salivary gland, binding NO, which is followed rapidly by the nitrosation of Cys-60 by a second NO and transition to ferrous (FeII+) heme (Weichsel et al., 2005b). An increased pH and low NO concentration in the host’s blood facilitates the reversal of this process and the release of large amounts of NO at the bite site with associated vasodilation. The nitrophorins then take on the role of antagonist, binding host histamine at their now ferric FeIII+ (Ribeiro and Walker, 1994). The remaining IPP5Ps all contain signal peptides and exonuclease-endonuclease-phosphatase (EEP) domains after their predicted cleavage sites. There is no published data concerning the role of these four IPP5Ps even though their expression levels are consistent, or higher than those of the well-studied nitrophorins. Structural models generated using the ExPASy SWISS-MODEL workspace seem to exclude them from coordinating with a heme ligand, which would at least suggest a different mechanism of NO binding, if they bind at all. The EPP domain suggests that they might act to modulate hemostatic states through breaking down polyphosphates and nucleotides at the bite site. The dominance and expansion of the IPP5Ps in the salivary gland of C. lectularius underscores the importance of their conserved structure in the physiology of hematophagy and warrants future study of their functionality.

            The two nudix hydrolase proteins enriched in the salivary gland, one of which has a signal sequence and the highest differential expression level of all the salivary transcripts (log2 FC = 20.08, p < 0.001), may be involved in anti-inflammatory and anti-immunity pathways during blood feeding. Vessels damaged during blood feeding expose collagen to von Willebrand factor, which mediates the binding and activation of platelets (Yun et al., 2016) triggering the release of inorganic phosphate polymers 60-100 units long. Anionic polymers promote the contact pathway (Gajsiewicz, Smith and Morrissey, 2017) of coagulation by strongly activating prekallikrein (factor XII) and kininogen (HK), which activate factor XI (Wiggins et al., 1977). A cascade of activated factors including factor XIa, Xa and Va ultimately leads to the production of thrombin and fibrin, which cycle through a positive feedback loop causing increased coagulation. Additionally, factor XIIa and HK link coagulation and inflammation by promoting the potent vasoactive mediator, bradykinin. The 23-amino acid Nudix hydrolase superfamily is considered a renaissance motif, conserved in 20,000 species and functioning in as disparate functions as metabolite degradation to ion channel gating (Mills et al., 2018). Interestingly, the most abundant salivary transcript is a Nudix hydrolase (LOC106666860) of 16.3 kDa which may be a previously identified factor X inhibitor (Valenzuela, Guimaraes and Ribeiro, 1996).  Additionally, extracellular nucleotides released from activated platelets, erythrocytes and endothelial cells regulate vascular tone, permeability and inflammation (Ralevic and Dunn, 2015). The dinucleotide uridine adenosine tetraphosphate (Up4A) causes vasoconstriction and its’ release is triggered by ATP, UTP and mechanical stress (Jankowski et al., 2005). The Nudix hydrolase in C. lectularius aligns to proteins with tetraphosphatase domains (Cordeiro et al., 2019) and may be responsible for inactivating Up4A. Odorant binding proteins are a protein family that is perhaps the most representative of rapid evolutionary expansion and divergence in C. lectularius. Amongst arthropods, the OBPs are archetypical of functional divergence (Robertson, 2019), exemplifying the point that any conclusion on protein functionality based solely on sequence similarity is only hypothetical. This applies to all the transcripts bioinformatically annotated here that have not been previously studied. Four out of ten of the most abundant salivary transcripts found in C. lectularius were either annotated as or have BLAST hits to OBPs. The most abundant OBP (log2 CPM = 14.97, p < 0.001) lacks a signal sequence and most likely plays a role in intracellular signal transduction, possibly related to host-seeking behavior. The secreted OBPs likely act as host immunomodulatory component antagonists, regulating hemostasis by binding histamine, serotonin, ATP and other important hemostatic molecules in the host (Andersen and Ribeiro, 2017). This is exemplified in the OBP-related D7 proteins in mosquitoes that bind serotonin, histamine and norepinephrine with high affinity (Calvo et al., 2006).

            There are four acetylcholinesterase-like salivary transcripts above 4-fold enrichment and at high abundance, with LOC106669436 and LOC106673686 having the 4th and 7th highest abundance of all salivary transcripts, respectively (log2 CPM = 14.97 and 13.86). The function of these esterase proteins remains unknown although they are conserved among blood-feeding arthropods. Aedes aegypti secretes several proteins with esterase activity that are of similar size (65 kDa) and align closely to the secreted Cimex acetylcholinesterase-like transcripts (Argentine and James, 1995). They also share conserved domain structure with platelet-activating factor (PAF)-acetylhydrolase proteins in the cat flea, Ctenocephalides felis (Cheeseman, Bates and Crampton, 2001). PAF has a very high affinity to G-protein coupled receptors on platelets, neutrophils and monocytes and may be responsible for cimicosis and eosinophil-mediated hypersensitivity reactions (Blom et al., 2019). A single apyrase present at high levels (log2 CPM = 14.54, p < 0.001) in the salivary gland shares substrates with the esterase family and was previously characterized in C. lectularius (Valenzuela et al., 1998). Enzymatically active apyrase proteins are often present in anthropophilic hematophagous arthropods (Cheeseman, 1998; Masoud et al., 2020) and humans have an apyrase closely related to that of the apyrase found in C. lectularius (Smith et al., 2002).

            Immunity within the bed bug salivary gland appears to be predominantly controlled by lysozyme-like proteins, which is consistent with that of other blood-feeding arthropods. A multigene family of lysozymes was identified in the initial bed bug sialome (Francischetti et al., 2010c) and is corroborated here by five closely related, highly enriched C-type lysozymes (Figure 4). Although lysozymes are not unique to hematophagous arthropods, a non-immune function related to the formation of the hemostatic plug in Triatoma pallidipennis (Noeske-Jungblut et al., 1995) suggests that antimicrobial peptides may function outside of their role in immunity. Lysozymes play a critical role in defending against systemic pathogen infection after blood feeding (Siva-Jothy et al., 2019) and it is possible that salivary gland-specific lysozymes act as the first line of defense by acting initially in the host.

            Of the top 1000 enriched salivary transcripts in C. lectularius, over 21% have signal peptides and are probably secreted. There has been little done in way of experimentation to functionally characterize these potentially important therapeutic proteins. As high-throughput sequencing grows increasingly economical and more arthropod sialomes are released, a demand for functional studies becomes necessary. Navigating these datasets and prioritizing potential targets will expediate the discovery of novel proteins with benefits to many scientific fields. Preliminary studies using transcriptomic data yield candidates as upregulated and expanded gene families that contain a signal peptide. Cimex lectularius remains an enduring anthropophilic ectoparasite whose evolutionary relationship with vertebrates represents an untapped pharmacopeia that demands closer examination.

References

Andersen, J. F. and Ribeiro, J. M. C. (2017) ‘Salivary kratagonists: scavengers of host physiological effectors during blood feeding’, Arthropod Vector: Controller of Disease Transmission, Volume 2. Academic Press, pp. 51–63. doi: 10.1016/B978-0-12-805360-7.00004-6.

Andrews, S. et al. (2010) ‘FastQC: a quality control tool for high throughput sequence data’.

Argentine, J. A. and James, A. A. (1995) ‘Characterization of a salivary gland-specific esterase in the vector mosquito, Aedes aegypti’, Insect Biochemistry and Molecular Biology. Pergamon, 25(5), pp. 621–630. doi: 10.1016/0965-1748(94)00103-O.

Assumpção, T. C. F. et al. (2008) ‘An insight into the sialome of the blood-sucking bug Triatoma infestans, a vector of Chagas’ disease’, Insect Biochemistry and Molecular Biology, 38(2), pp. 213–232. doi: 10.1016/j.ibmb.2007.11.001.

Atyame, C. M. et al. (2019) ‘Insecticide resistance genes affect Culex quinquefasciatus vector competence for West Nile virus’, Proceedings of the Royal Society B: Biological Sciences. Royal Society Publishing, 286(1894). doi: 10.1098/rspb.2018.2273.

Baptist, B. A. (1941) ‘The morphology and physiology of the salivary glands of Hemiptera-Heteroptera’, Journal of Cell Science. The Company of Biologists, s2-83(329), pp. 91–139. doi: 10.1242/jcs.s2-83.329.91.

Berridge, M. J. and Patel, N. G. (1968) ‘Insect salivary glands: stimulation of fluid secretion by 5-hydroxytryptamine and adenosine-3’,5’-monophosphate.’, Science. American Association for the Advancement of Science, 162(852), pp. 462–463. doi: 10.1126/science.162.3852.462.

Blom, M. et al. (2019) ‘Eosinophils and platelet-activating factor’, in Gleich, G. (ed.) Eosinophils in Allergy and Inflammation. CRC Press, pp. 162–166.

Bolger, A. M., Lohse, M. and Usadel, B. (2014) ‘Trimmomatic: a flexible trimmer for Illumina sequence data’, Bioinformatics. Oxford University Press, 30(15), pp. 2114–2120. doi: 10.1093/bioinformatics/btu170.

Calvo, E. et al. (2006) ‘Function and evolution of a mosquito salivary protein family’, Journal of Biological Chemistry, 281(4), pp. 1935–1942. doi: 10.1074/jbc.M510359200.

Cheeseman, M. T. (1998) ‘Characterization of apyrase activity from the salivary glands of the cat flea Ctenocephalides felis’, Insect Biochemistry and Molecular Biology, 28(12), pp. 1025–1030. doi: 10.1016/S0965-1748(98)00093-9.

Cheeseman, M. T., Bates, P. A. and Crampton, J. M. (2001) ‘Preliminary characterization of esterase and platelet-activating factor (PAF)-acetylhydrolase activities from cat flea (Ctenocephalides felis) salivary glands’, Insect Biochemistry and Molecular Biology. Elsevier Ltd, 31(2), pp. 157–164. doi: 10.1016/S0965-1748(00)00113-2.

Chen, Y. et al. (2020) ‘An aphid RNA transcript migrates systemically within plants and is a virulence factor’, Proceedings of the National Academy of Sciences of the United States of America. National Academy of Sciences, 117(23), pp. 12763–12771. doi: 10.1073/pnas.1918410117.

Cordeiro, C. D. et al. (2019) ‘NUDIX hydrolases with inorganic polyphosphate exo- and endopolyphosphatase activities in the glycosome, cytosol and nucleus of Trypanosoma brucei.’, Bioscience reports. Portland Press Limited, 39(5), p. BSR20190894. doi: 10.1042/BSR20190894.

Francischetti, I. M. B. et al. (2002) ‘Toward a catalog for the transcripts and proteins (sialome) from the salivary gland of the malaria vector Anopheles gambiae.’, The Journal of experimental biology, 205(Pt 16), pp. 2429–51.

Francischetti, I. M. B. et al. (2010) ‘Insight into the sialome of the bed bug, Cimex lectularius.’, Journal of proteome research. NIH Public Access, 9(8), pp. 3820–31. doi: 10.1021/pr1000169.

Gajsiewicz, J. M., Smith, S. A. and Morrissey, J. H. (2017) ‘Polyphosphate and RNA differentially modulate the contact pathway of blood clotting.’, The Journal of biological chemistry. American Society for Biochemistry and Molecular Biology, 292(5), pp. 1808–1814. doi: 10.1074/jbc.M116.754325.

Grimaldi, D. and Engel, M. S. (2005) ‘Evolution of the Insects’. Cambridge University Press, Cambridge, England.

Harris, S. J. et al. (2008) ‘Phosphoinositide lipid phosphatases: natural regulators of phosphoinositide 3-kinase signaling in T Lymphocytes’, Journal of Biological Chemistry, 283(5), pp. 2465–2469. doi: 10.1074/jbc.R700044200.

Jankowski, V. et al. (2005) ‘Uridine adenosine tetraphosphate: a novel endothelium- derived vasoconstrictive factor’, Nature Medicine, 11(2), pp. 223–227. doi: 10.1038/nm1188.

Kim, D., Langmead, B. and Salzberg, S. L. (2015) ‘HISAT: a fast spliced aligner with low memory requirements’, Nature Methods. Nature Publishing Group, 12(4), pp. 357–360. doi: 10.1038/nmeth.3317.

Li, H. et al. (2009) ‘The Sequence alignment/map format and SAMtools’, Bioinformatics, 25(16), pp. 2078–2079. doi: 10.1093/bioinformatics/btp352.

Masoud, H. M. M. et al. (2020) ‘Apyrase with anti-platelet aggregation activity from the nymph of the camel tick Hyalomma dromedarii’, Experimental and Applied Acarology. Springer, 80(3), pp. 349–361. doi: 10.1007/s10493-020-00471-9.

Mills, J. L. et al. (2018) ‘Functional characterization of NUDIX hydrolases’, The FASEB Journal. Federation of American Societies for Experimental Biology, 32(1_supplement), pp. 796.16-796.16. doi: 10.1096/fasebj.2018.32.1_supplement.796.16.

Noeske-Jungblut, C. et al. (1995) ‘Triabin, a highly potent exosite inhibitor of thrombin’, Journal of Biological Chemistry. American Society for Biochemistry and Molecular Biology, 270(48), pp. 28629–28634. doi: 10.1074/jbc.270.48.28629.

Omori, N. (1941) ‘Comparative studies on the ecology and physiology of common and tropical bed bugs, with special reference to the reactions to temperature and moisture.’, Journal of the Medical Association of Formosa, 40(3), pp. 555–636.

Pertea, M. et al. (2016) ‘Transcript-level expression analysis of RNA-seq experiments with HISAT, StringTie and Ballgown’, Nature Protocols, 11(9), pp. 1650–1667. doi: 10.1038/nprot.2016.095.

Peta, V. and Pietri, J. E. (2021) ‘Experimental infection of bed bugs (Cimex lectularius L.) with Burkholderia multivorans’, Medical and Veterinary Entomology. Blackwell Publishing Ltd. doi: 10.1111/mve.12520.

Ralevic, V. and Dunn, W. R. (2015) ‘Purinergic transmission in blood vessels’, Autonomic Neuroscience, 191, pp. 48–66. doi: 10.1016/j.autneu.2015.04.007.

Ribeiro, J. M. C. et al. (2004) ‘Exploring the sialome of the blood-sucking bug Rhodnius prolixus.’, Insect biochemistry and molecular biology, 34(1), pp. 61–79.

Ribeiro, J. M. C. and Arcà, B. (2009) ‘From sialomes to the sialoverse. an insight into salivary potion of blood-feeding insects.’, in Advances in Insect Physiology. Academic Press Inc., pp. 59–118. doi: 10.1016/S0065-2806(09)37002-2.

Ribeiro, J. M. and Walker, F. A. (1994) ‘High affinity histamine-binding and antihistaminic activity of the salivary nitric oxide-carrying heme protein (nitrophorin) of Rhodnius prolixus’, Journal of Experimental Medicine, 180(6), pp. 2251–2257. doi: 10.1084/jem.180.6.2251.

Robertson, H. M. (2019) ‘Molecular evolution of the major arthropod chemoreceptor gene families’, Annual Review of Entomology. Annual Reviews, 64(1), pp. 227–242. doi: 10.1146/annurev-ento-020117-043322.

Robinson, M. D., McCarthy, D. J. and Smyth, G. K. (2010) ‘edgeR: a Bioconductor package for differential expression analysis of digital gene expression data’, Bioinformatics, 26(1), pp. 139–140. doi: 10.1093/bioinformatics/btp616.

Rosenfeld, Jeffrey A et al. (2016) ‘Genome assembly and geospatial phylogenomics of the bed bug Cimex lectularius.’, Nature communications. Nature Publishing Group, 7, p. 10164. doi: 10.1038/ncomms10164.

Santos, A. et al. (2007) ‘The sialotranscriptome of the blood-sucking bug Triatoma brasiliensis (Hemiptera, Triatominae)’, Insect Biochemistry and Molecular Biology, 37(7), pp. 702–712. doi: 10.1016/j.ibmb.2007.04.004.

Siva-Jothy, M. T., Zhong, W., Naylor, R., Heaton, L., Hentley, W., & Harney, E. (2019) ‘Female bed bugs (Cimex lectularius L) anticipate the immunological consequences of traumatic insemination via feeding cues’. PNAS, 116(1), pp. 14682–14687. doi: 10.1073/pnas.1904539116.

Smith, T. M. et al. (2002) ‘Cloning, expression, and characterization of a soluble calcium-activated nucleotidase, a human enzyme belonging to a new family of extracellular nucleotidases’, Archives of Biochemistry and Biophysics. Academic Press, 406(1), pp. 105–115. doi: 10.1016/S0003-9861(02)00420-4.

Stavrou, B. M. (2003) ‘Diadenosine polyphosphates: postulated mechanisms mediating the cardiac effects.’, Current medicinal chemistry. Cardiovascular and hematological agents. Curr Med Chem Cardiovasc Hematol Agents, 1(2), pp. 151–169. doi: 10.2174/1568016033477513.

Talbot, B. et al. (2017) ‘Host association and selection on salivary protein genes in bed bugs and related blood-feeding ectoparasites’, Royal Society Open Science. The Royal Society, 4(6), p. 170446. doi: 10.1098/RSOS.170446.

Valenzuela, J. G. et al. (1998) ‘Purification, cloning, and expression of an apyrase from the bed bug Cimex lectularius: A new type of nucleotide-binding enzyme’, Journal of Biological Chemistry, 273(46), pp. 30583–30590. doi: 10.1074/jbc.273.46.30583.

Valenzuela, J. G., Chuffe, O. M. and Ribeiro, J. C. (1996) ‘Apyrase and anti-platelet activities from the salivary glands of the bed bug Cimex lectularius’, Insect Biochemistry and Molecular Biology. Pergamon, 26(6), pp. 557–562. doi: 10.1016/S0965-1748(96)00041-0.

Valenzuela, J. G., Guimaraes, J. A. and Ribeiro, J. M. C. (1996) ‘A novel inhibitor of factor X activation from the salivary glands of the bedbug Cimex lectularius’, Experimental Parasitology, 83(2), pp. 184–190. doi: 10.1006/expr.1996.0065.

Valenzuela, J. G., Walker, F. A. and Ribeiro, J. M. (1995) ‘A salivary nitrophorin (nitric-oxide-carrying hemoprotein) in the bedbug Cimex lectularius’, The Journal of Experimental Biology, 198(7), pp. 1519–1526.

Wang, J. et al. (2016) ‘Evolutionary fates and dynamic functionalization of young duplicate genes in Arabidopsis genomes.’, Plant physiology. American Society of Plant Biologists, 172(1), pp. 427–40. doi: 10.1104/pp.16.01177.

Weichsel, A. et al. (2005) ‘Heme-assisted S-nitrosation of a proximal thiolate in a nitric oxide transport protein’, Proceedings of the National Academy of Sciences, 102(3), pp. 594–599. doi: 10.1073/pnas.0406549102.

Wiggins, R. C. et al. (1977) Role of high-molecular-weight kininogen in surface-binding and activation of coagulation Factor XI and prekallikrein, Medical Sciences.

Yun, S.-H. et al. (2016) ‘Platelet activation: the mechanisms and potential biomarkers.’, BioMed research international. Hindawi Limited, 2016, p. 9060143. doi: 10.1155/2016/9060143.

Zhang, Y. et al. (2020) ‘Depicting the RNA virome of hematophagous arthropods from Belgrade, Serbia’, Viruses. MDPI AG, 12(9), p. 975. doi: 10.3390/v12090975.

CHAPTER II 

dsRNA-MEDIATED KNOCK-DOWN OF NITROPHORIN IN CIMEX LECTULARIUS AND ITS IMPACT ON BITE REACTIONS IN HUMANS

Background

            Cimex lectularius is an ectoparasite that is genetically and morphologically divided into 2 different lineages that prefer to feed on either humans, bats or swallows (Balvín et al., 2012). While they both exist in the same geographic regions, mitochondrial and microsatellite genetic data indicate that there is no contemporary interbreeding or gene flow occurring between the lineages (Booth et al., 2015), which diverged ~245,000 years ago. The lineage of C. lectularius that feeds on humans, often cohabitates with their host and has made a dramatic resurgence as a pest species, especially in developing countries (Doggett et al., 2012). Defining the relationship between the parasite and host extends beyond genetic background and geographic distribution and has important immunological and clinical consequences. The unique and persistent relationship between humans and bed bugs is associated with an acute inflammatory reaction called cimicosis, which causes a dermal reaction on the afflicted patient. The course of the reaction varies based on previous exposure and immunological background and can be delayed by a week, an hour or totally absent depending on the individual and the reaction mechanism and pathophysiology remains unclear. If we define a consistent reaction in affected patients, marked by a specific immune cell and effector profile, we can gain insight into vertebrate immunology and develop treatments and preventative measures to combat cimicosis.

            Cimex lectularius is a robust parasite, feeding every few days depending on host availability (Reinhardt, Isaac and Naylor, 2010). This means that a human host could be bitten thousands of times over the course of an infestation. The initial reaction experienced by the human host fed upon for the first time by a bed bug usually appears after a week (Reinhardt et al., 2009) but occurs more rapidly upon repeated feedings (Goddard, Edwards and de Shazo, 2011). Some people appear to be immune with one survey of emergency room patients who reported having past or present bed bug infestations indicating that many did not develop the signature post-bed bug feeding pruritic rash (Sheele et al., 2019), although the exact number is difficult to determine.

            The antigenic determinant of the inflammatory allergic reaction is most likely contained within the salivary glands, as bed bugs lacking these organs did not elicit a response in a human volunteer (Goddard and Edwards, 2013). In this study, a group of bed bugs had their salivary glands removed and were still able to feed. The bites from intact, control bed bugs elicited an itchy erythematous lesion in 2 to 5 days, while those without did not. Interestingly, the volunteer indicated feeling a “biting” sensation from the experimental bed bugs who lacked salivary glands, suggesting that the saliva contains anesthetic compounds. A 2006 case study implicated Cimex nitrophorin as the allergic determinant as it was targeted by the allergy-mediating antibody IgE in a sensitive patient’s serum (Leverkus et al., 2006). The patient was a 43-year-old female who reported staying at a hotel twice, separated by a year. After the first visit she noticed small hemorrhagic macules that healed quickly but upon the second visit she saw multiple erythematous papules, nodules, and blisters that only resolved with the application of clobetasol propionate 0.5% (Leverkus et al., 2006). An immunoblot using the patients serum run against a salivary gland solution showed strong binding of anti-IgE secondary antibodies at approximately 32 kDa, which is where nitrophorin comes up on PAGE/MS/MS using salivary extracts (Francischetti et al., 2010c). There was no IgE binding using a healthy individual’s serum and the IgG binding patterns were the same between the two subjects. IgE mediates type I hypersensitivity reactions (Sutton et al., 2019) and can trigger life-threatening anaphylaxis in sensitized individuals upon cross-linking with soluble multivalent allergens when bound to FcεRI on effector cells (Schwartz, 2004). Another study of 11 human subjects showed no consistent IgG response in subjects exposed to bed bug bites (Sheele et al., 2020). In this study, patients were bitten weekly by 3 male bed bugs for 1 month after which their serum was run on an immunoblot against salivary extracts. Anti-IgG secondary antibodies indicated an increased reaction to a 12 to 13 kDa protein in 6 patients and to 30, 40 and 70 kDa proteins in 5 patients. The most likely effector triggering the 40 kDa IgG response was the salivary apyrase, but the results were inconclusive and lacked data on IgE binding (Sheele et al., 2020). Similarly inconclusive results were seen in cytokine and antibody levels of patients undergoing the same experimental feeding regime (Sheele et al., 2017). They found that there was no significant change in total IgG, IgG1, IgG2, IgG4, or IgE levels from the 1st to the 4th feeding and that there was no significant change in serum TNFα, IL-1 β, IL-4, IL-5, IL-6, IL-10, IFNγ, and IL-17A levels before and one hour after feeding. They did find an increase in IgG3 levels after the 4th feeding and that lower IL-6 levels after feeding were associated with increased pruritis (Sheele et al., 2017).

Material and methods

2.1.1             Maintenance and storage of experimental colony arthropods

            Control or experimental individuals of the Harlan strain of C. lectularius were maintained in glass screw top vials 3 cm by 7 cm by 2 cm in size. Small cardboard strips were folded and cut to provide shelter for the bed bugs. Vials were grouped and placed in 9 cm by 15 cm by 6 cm plastic pipette boxes to facilitate transport between housing and injection facilities. Bed bugs use for experimental injections and qPCR were reared from the first nymphal stage to adulthood to synchronize age. The first nymphal stage was identified by the absence of blood in the abdomen, which is clearly visible through the transparent cuticle. A fine paint brush was used to separate the first stage nymphs from colony to new jars where they were blood fed and maintained at 30°C to maximize growth to adulthood (Omori, 1941).

2.1.2             Production of dsRNA

            Total genomic DNA from a single female individual of C. lectularius was extracted for use as the PCR template. T7 templates used by an RNA polymerase must have the appropriate adapters, not present in the endogenous DNA, for the polymerization to yield ultra-high levels of dsRNA on the 400-650 moles per every mole of DNA template. The promoter sequence used for the MEGAscript™ T7 Transcription Kit was affixed 5’ upstream of the transcribed region and contained the promoter sequence: 5’- GAATTAATACGACTCACTATAGGG-3’ followed by the Cimex specific primer sequence, which will be in the final dsRNA. The promoter region must be on both the forward and the reverse primers, with an identical 5’ to 3’ sequence on both primers to obtain double-stranded RNA. Cimex specific primers were designed using Primer3 with settings designating a 500 base pair product (Kõressaar et al., 2018).

            The template was amplified using Q5® High-Fidelity DNA Polymerase (M0491) and Nitro_531 primers (Appendix) at final concentrations of 1x Q5 reaction buffer, 200 uM dNTPs, 0.5 uM Nitro_531 forward and reverse primers, 0.3 µl (28 ng) C. lectularius genomic DNA and 0.02 U/µl of Q5 High-fidelity DNA Polymerase in 0.2 ml PCR tubes under the following reaction conditions: 1 cycle at 98°C for 30 sec, 35 cycles each at 98°C for 10 sec, 66°C for 30 sec and 72°C for 20 sec, and 1 cycle at 72°C for 2 min and a final hold at 4°C. A 1.0% TAE gel was cast using 0.5 g Denville Scientific Inc agarose (Cat #gr140-500) and 50 ml of 10x TAE buffer heated for 3 min on high in a GE microwave oven. The resulting amplified PCR products were run on a 14 cm long Thermo Scientific, Owl Easycast B1A gel rig connected to a Bio Rad PowerPac Basic 300 V, 75 W power supply run at 90 V for 40 min. The resulting raw PCR amplicons were cleaned of residual protein, primers and buffer using QIAquick Spin columns and the QIAquick PCR Purification Kit For purification of PCR products (100 bp – 10 kb) protocol. Briefly, 200 µl of total raw PCR product was combined with 1,000 µl of Buffer PB and gently mixed by pipetting the mixture up and down. The pH was adjusted using 3 M sodium acetate until the color turned yellow, indicating an acidic pH. This was added to QIAquick spin column in a 2 ml collection tube and centrifuged at 16,000 g for 1 min. The flow through was discarded and the column was returned to the collection tube. The column was then washed with 750 µl of buffer PE, which was centrifuged at 16,000 g for 1 min. The flow through was discarded and the column was dried by centrifugation at 16,000 g for 1 min. The QIAquick column was placed in a clean 1.5 ml microcentrifuge tube and the purified PCR product was eluted with 30 µl of water (pH 7.0) heated to 55°C by adding the water directly to the column and incubating at room temperature for 1 min followed by centrifugation at 16,000 g for 1 min. The purified dsNitro_531 fragment was quantified using the Thermo Scientific Nanodrop One-C Microvolume UV Spectrophotometer.

            Double-stranded RNA was transcribed using the MEGAscript™ T7 Transcription Kit. The 10x Reaction Buffer for the transcription kit contains spermidine and must be kept at room temperature during the reaction assembly to avoid co-precipitation of the DNA template. Two µl ATP, 2 µl CTP, 2 µl GTP, 2 µl UTP 2ul 10x reaction buffer, 8 µl 125 ng/µl purified Nitro_531 DNA template and 2 µl of enzyme mix were added to a 0.2 ml nuclease-free PCR tube at room temperature. The reaction mix was mixed by pipetting up and down and pulse centrifugation. The reaction mix was incubated 37°C for 4 hr before removing the reaction from the thermocycler, 1 µl of TURBO DNase was mixed into the reaction by pipetting and the reaction proceeded for an additional 15 min at 37°C. The final product was brought to 50 µl through the addition of 29 µl of nuclease-free water and cleaned using the Monarch® RNA Cleanup Kit (Cat #T2040L). The dsRNA product is mixed with 100 µl RNA Cleanup Binding Buffer in the original 0.2 ml PCR reaction tube. The binding buffer/dsRNA mixture was added to 150 µl of 200 proof, molecular biology grade absolute ethanol (Cat #T038181000) in a new 1.5 ml microcentrifuge tube. All of this was added to a spin column in a collection tube and the tube was centrifuged at 16,000 g for 1 min at room temperature. The flow through was discarded and the column was put back into the original collection tube. The wash was carried out by applying 500 µl of RNA Cleanup Wash Buffer to the column and centrifuging at 16,000 g for 1 min. The was repeated and the column was placed into a clean RNase-free 1.5 ml microcentrifuge tube. No incubation time was needed for the T2040 kit so 100 µl nuclease-free water was added directly to the column and the tube was centrifuged at 16,000 g for 1 min. The 100 µl of eluted dsRNA was quantified immediately using the Nanodrop One-C and the reaction tube was stored at -80°C until injection.

2.1.3             Injection of bed bugs

Bed bugs were injected using 3 in borosilicate glass capillary tubes with a 1 mm outer diameter (World Precision Instruments; Cat # 1B100-3). The capillary tubes were pulled using a Narishige PC-10 capillary needle puller. The bottom adapter was loaded with all 4 weights and temperature setting 2 was set to 50.4°C. The capillary needle was loaded so that the end extending out of the top of the device reached about 2 cm above the machine and both ends were firmly affixed by twisting the plastic knobs clockwise. Care was taken to not turn the knobs too tightly as to avoid breaking the glass. The knob on the bottom of the device was turned to “step1” and the plastic door affixed to the heating coil was closed. When the weight dropped both pulled capillary needles were carefully removed and placed in a square petri dish containing a1” and the plastic door affixed to the heating coil was closed. When the weight dropped both pulled capillary needles were carefully removed and placed in a square petri dish containing a piece of double-sided tape. The tip of the piece of double-sided tape. The tip of the needle was broken under the microscope

Figure 2.1           The injection device used for mouth aspiration of C. lectularius. (a) The aspirator has a red mouthpiece which is enclosed in a 2 ml microcentrifuge tube here to promote hygiene and prevent cross-contamination. The red circle encompasses the adapter, which holds the borosilicate glass capillary needle. (b) The borosilicate glass capillary needle pulled using the PC-10 Narishige needle puller set to 50.4°C is manually delineated every 3.84 mm to indicate 1 ul. This facilitates injection, which is manually observed between each mark, making sure to not evacuate the needle past the beveled tip as the volume there can vary. (c) The entire apparatus with the borosilicate capillary needle affixed to the adapter outlined with a red circle. This particular image displays the manipulation of the specimen, which is not affixed to a microscope slide or double-sided tape.

so that it could be filled using capillary action. This was done by gently sliding the tip of the capillary needle over the edge of a pair of forceps until just the very tip breaks off. The capillary needles hold 1 µl in every 3.79 mm length of the unpulled needle, so this length was measured and marked on the glass with a black sharpie marker. The capillary needle was attached to the end of the clear adapter on the aspirator and was loaded by using the red mouthpiece to gently inhale 1 µl of dsRNA Baptofect reagent for each individual injection. The bed bugs were taken individually from their vials and affixed to a strip of double-sided tape under the stereoscope using a paint brush. Their ventral side was facing up and they were injected within the 6th to 10th parastergite segments with 1 µl of injection mixture.

2.1.4             Quantification of gene expression in C. lectularius using qPCR

            The synthesis of cDNA template was done using the Thermo Scientific RevertAid First Strand cDNA Synthesis Kit (Cat #K16621). RNA was first treated with DNase to remove genomic DNA carried over from the RNA extraction. Reactions were brought up to 10 µl with nuclease-free water in 200 µl PCR tubes and contained 1 ug of RNA, 1 µl of 10x Reaction Buffer with MgCl2 and 1 µl of DNaseI, RNase free (Cat #EN0521). The reaction was incubated at 37°C for 30 min followed by the addition of 1 µl 50 mM EDTA and a 65°C incubation for 10 min. All first strand reactions contained 1 ng to 500 ng of total RNA depending on the concentration of the input template. First strand cDNA synthesis reactions were mixed in a 200 µl nuclease-free PCR tube and contained 1 – 500 ng of total RNA template and 1 µl of oligo (dT)18 primers which was brought to 12 µl with nuclease-free water and incubated at 65°C for 5 min before immediately chilling on ice and pulse centrifuging. The reaction was then brought to 20 µl with 4 µl of 5x reaction buffer, 1 µl of RiboLock RNase Inhibitor, 10 mM dNTP mix and RevertAid M-MuLV RT before mixing with a pipette and pulse centrifugation. The reaction was then incubated at 42°C for 60 min followed by a 5 min 70°C reaction termination step. All cDNA samples were transferred to a 1.5 ml nuclease-free microcentrifuge tube, diluted with 30 µl of nuclease-free water and stored at 80°C until needed.

            Quantitative, real-time PCR reaction were done using 5x HOT FIREPol ® EvaGreen ® qPCR Mix Plus (SolisBioDyne; Cat #08-25-00001). The master mix was assembled on ice in a 1.5 ml nuclease-free microcentrifuge tube. The final concentrations in the reactions mix were 1x HOT FIREPol ® EvaGreen® qPCR Mix Plus, 100 nM forward and reverse primer and 1  µl  of cDNA template. The master mix was assembled without template or primer (just water and 1x HOT FIREPol ® EvaGreen® qPCR Mix Plus) and 18 µl was distributed to each of the 200 µl MicroAmp Fast Reaction Tubes (Applies Biosystems). The primers and templates were then individually loaded, 1 µl at a time to each reaction tube and they were firmly sealed with Optical 8-Cap Strips (Applied Biosystems). Care was taken to prevent marking the tubes in any way including with marker labels, so keeping the tubes oriented in the correct configuration was vital. The tubes were loaded into a 96-well, 200 µl tube rack and any bubbles were removed by carefully flicking the bottom of the tube using a finger. The tubes were then pulse centrifuged in and loaded onto the qPCR machine.

            The Applied Biosystems QuantStudio 3 machine was used for real-time PCR. The machine must be powered on to load the device, which is done by pushing the eject button on the top-right corner of the touch screen. Using the QuantStudio Design and Analysis Software v1.5.1 run on the Dell laptop connected to the qPCR machine, the run properties were entered. After executing the software, “create new experiment” was selected and the name of the experiment was entered in the top experimental properties field. The “96-Well 0.1-mL Block” was chosen and must be selected for this device, or the machine will delete any created template upon starting the run. The experimental type was either “Standard Curve” to validate any unpublished primers or “Comparative Ct (ΔΔCT)” for experimental runs. The chemistry was “SYBR Green Reagents” for all experiments run here and the run mode was “Standard”. The “Method” option was chosen from the overlay menu at the top of the window and the method was entered according to the specific chemistry and reagents used. For 5x HOT FIREPol EvaGreen qPCR Mix Plus the qPCR cycles were: 1 initial denaturation and hot-start polymerase activation at 95°C for 15 min, 40 cycles of 95°C for 15 sec, 54°C for 20 sec and 72°C for 20 sec, and the final melt curve stage of ramp to 95°C for 15 sec by 1.6°C/sec, ramp to 60°C for 1 min by 1.6°C/sec and ramp to 95°C for 15 sec by 0.1 °C/sec to record the disassociation of the amplicons.

2.1.5             Cimex lectularius bite assays in a human volunteer

            All bite assays were done using a single human volunteer and protocol approved by the institutional review board (IRB #17-363). PBS-injected and experimental bed bugs were stored in individual vials and were maintained at 27°C to recover from injections for 24 hrs. After recovery, both groups of bed bugs were allowed to probe on a human volunteer to evacuate their salivary glands without being allowed to take a blood meal. Individual bed bugs were placed on the back of the hand one at a time where they would move their proboscis up and down into the human volunteer’s skin before becoming stationary. From the time they stopped moving their proboscis, 3 – 5 sec was measured, and a pair of forceps was gently brushed under their mouthparts to stop the feeding process. This was repeated 5 times, making sure that no blood was consumed during the feeding process. After draining their salivary glands, the bed bugs were returned to their designated vials and incubated at 27°C until the initial feeds were performed. At 48 hrs post injection, the first blood feeds were performed on the human volunteer. Each bed bug was fed individually to record all variables accurately, starting at the base of the forearm near the wrist with palms facing up. The bed bugs were placed in vials labeled with tape, so that the individual taking the measurements is unaware of whether they are recording the experimental or control groups. The bed bugs were placed on the human volunteer’s wrist, and the time from initial contact with the skin to probing with the proboscis is recorded. Any time the proboscis was removed the time was recorded until the bed bugs were replete, at which point the total time was recorded. Weights of each bed bug was taken before and after the blood meal using a Torbal analytical balance (Cat #AGZN120) by placing the individuals on a weigh boat prior to recording with both balance doors closed. The bite site reactions were allowed to develop for 6 hrs prior to recording the size of inflammation. Stereoscope photos of each bite were taken with the Zeiss Discovery.V12 with the magnification set to 8.1x in a field of view of 32.0 mm. Two Zeiss CL 6000 LED lights in metal armatures (Cat #435441) were affixed 6 cm away from the bite epicenter, set at a light level of 80. The Zeiss Axiocam 208 color was used to take the photos with the auto exposure level set to 73.

            ImageJ was run in the Fiji environment to quantify the area of the bite reaction. First, the ImageJ executable was opened to run the software. The file photo for each bite was opened using “file” dropdown and clicking on “open”. The image was converted to 8-bit by selecting the “image” dropdown icon and the “type” and “8-bit” selections. The “straight” line icon was select from the ImageJ toolbar and was used to extend a line across the 1 mm scale bar that was included in each bite photo taken with the stereoscope. The “analyze” dropdown was opened and the “set scale” menu was selected. The “known distance” and “unit of length” queries were changed to “1” and “mm”, respectively, and the “global” box was checked. The “image” dropdown was opened, followed by the “adjust” and “threshold” options to bring up the scalar adjustments for the photo’s 8-bit color display. The levels were adjusted to encompass the secondary and primary inflammation in the photo with a red color. It is at this point where subjective bias can be introduced into the experiment, so it is important that a single individual, unaware of the experimental variables being measured, make all the determinations to be included in the threshold setting. Once the bite area is determined, the wand tracing tool was selected and used to ctrl+left click the red area. The “analyze” dropdown and “tools” then “ROI manager” was selected, and the “add” button was pushed from the popup menu. The area was selected, and the “measure” button was pushed to give the mean measurement of the selected area. The lighter primary and darker secondary inflammation were measured for all photos at 6 hr, 24 hr and 48 hr post bite for the experimental and control groups.

Figure 2.2           An example of a single photo analyzed under different brightness settings in ImageJ. The image was captured using the Zeiss Discovery.V12 with the magnification set to 8.1x in a field of view of 32.0 mm. Two Zeiss CL 6000 LED lights in metal armatures were affixed 6 cm away from the bite epicenter, set at a light level of 80. The Zeiss Axiocam 208 color was used to take the photos with the auto exposure level set to 73. (a) The unaltered photo lacks the contrast needed to delineate the immunological junction erythema and unaffected skin. (b) In the adjusted photo, contrast is brought up to the point where brighter areas in the photo turn white, at which point the brightness is dropped to clearly show inflammatory delineation. (c) The primary erythema is outlined in yellow and will always enclose secondary erythema. The outline here was drawn with Inkscape and does not represent the outlines described in the protocol or available in ImageJ. (d) Secondary inflammation is characterized as a more intense erythema which can be differentiated from the initial reaction. Because of its intensity, it is easier to objectively measure.

Results

2.1.6             Nitrophorin expression under standard growth conditions

            All primer sets used for the 6 gene nitrophorin panel were validated by qPCR using a 5-fold serial dilution of purified PCR fragments. Each primer set had R2 and efficiency values from 0.95-0.99 and 90-105%, respectively. Expression profiles from nymphal bed bugs were taken from pooled individuals in groups of 3 for each stage. Average ΔΔCT values indicate relative expression compared to validated beta-tubulin and UBC housekeeping genes averaged together. In stage 1 nymphs, the ΔΔCT of LOC106663254 was 1.82, in LOC106662975 the ΔΔCT was 1.07, ΔΔCT in LOC106662976 was 11.88, ΔΔCT in LOC106662977 was 8.17, ΔΔCT in LOC106662978 was 1.89 and ΔΔCT in LOC106662979 was 1.67 (figure 2.4). The patterns of relative expression values were similar across nymphal stage for all nitrophorin and nitrophorin-like genes (figure 2.5).

            Expression profiles of nitrophorin-76 and nitrophorin-77 for adult bed bugs reared from eggs averaging 53 days old are seen in figure 2.5. At 13 days starved adult bed bugs averaged a ΔΔCT value of 75. At 14 days starved this value increased to 452,227. Two days after blood feeding the ΔΔCT was 13.23, at 3 days it was 2.74 and at 6 days it was 0.47.

Figure 2.3         Gene expression analysis of nitrophorin and nitrophorin-related genes in 1st stage nymphs of C. lectularius. (a) Stereoscope image depicting a 1st stage nymph of C. lectularius with no blood meal. Salivary glands are indicated by red arrows and have a pinkish-orange tint. (b) Gene expression analysis measured using the Applied Biosystems QuantStudio 3 machine. Each gene displayed on the x-axis is labeled by the final 2 numbers of its LOC designation on NCBI. Relative expression levels are depicted on the y-axis and are designated as ΔΔCt. All measurements for the N1 stage were made from cDNA samples obtained from 3 biological replicates of pooled nymphs, each containing 10 individuals who were newly emerged no more than 10 days prior to RNA extraction.

Figure 2.4           Gene expression analysis of the nitrophorin and nitrophorin-related genes in C. lectularius. All data was measured using the Applied Biosystems QuantStudio 3 machine. Each gene displayed on the x-axis is labeled by the final 2 numbers of its LOC designation on NCBI. Relative expression levels are depicted on the y-axis and are designated as ΔΔCt. (a) Stage 2 nymphs as measured from 3 biological replicates containing 10 pooled nymphs each. (b) Stage 3 nymphs as measured from 3 biological replicates containing 10 pooled nymphs each. (c) Stage 4 nymphs as measured from 3 biological replicates containing 10 pooled nymphs each. (d) Stage 5 nymphs as measured from 3 biological replicates containing 10 pooled nymphs each.

Figure 2.5           Gene expression analysis of nitrophorin-76 and nitrophorin-77 in synchronized adult bed bugs reared from eggs before and after a blood meal. Bed bugs were brought to adulthood in 40 days and blood fed on the 14th day after that. This graph shows the 2 days prior to that blood meal and 6 days after. Each day contains 3 adult bed bugs representing 3 biological replicates per timepoint.

2.1.7             dsRNA production

            PCR cleanups of the 76_CiT7 and 77_CiT7 gave DNA yields of 128 and 315 ng/µl in 50 µl of nuclease-free water. Both were run on an agarose gel to confirm their size and submitted for Sanger sequencing to confirm their identity (figure 2.6). These templates initially gave dsRNA yields of 615 and 408 ng/µl of dsRNA for 76_CiT7 and 77_CiT7, respectively, in 50 µl of nuclease-free water using the Megascript kit. These reactions were concentrated to 1,252 and 1,341 ng/µl for 76_CiT7 and 77_CiT7, respectively, for injection. All reactions for the initial nitro-76 and nitro-77 injections were diluted down to 1,000 ng/µl immediately prior to abdominal injection into individual bed bugs.

            Total genomic DNA extracted from a single individual of C. lectularius yielded 83.3 ng/µl by Nanodrop with 260/280 and 260/230 values of 1.86 and 1.98, respectively. The raw Nitro_531 product ran between 400 and 500 base pairs on an agarose gel (figure 2.7; c). The cleaned-up fragment run through the QIAquick spin column yielded 30 µl at 125 ng/µl with 260/280 and 260/230 values of 1.81 and 2.10, respectively. The Megascript kit used for dsRNA synthesis using the Nitro_531_T7 template yielded 1,960 ng/µl in 50 ul, which was diluted to 1,000 ng/µl with 48 µl of Baptofect transfection reagent. This was the final solution used for injection.

Figure 2.6           The two highly expressed nitrophorin genes in C. lectularius, nitro-76 and nitro-77. (a) Screenshots from the NCBI gene maps for each of the two genes. On the top is LOC106662976 (nitro-76) and on the bottom is LOC106662976 (nitro-77). The blue and red selected regions flank areas targeted by dsRNA and amplified in subsequent panels. (b) The agarose gel showing the PCR-amplified regions for each of the nitro-76 and nitro-77 T7 dsRNA templates. The top and bottom red outlined arrows point to the 602 and 535 bp fragments, respectively. (c) An example of the chromatogram results from the Sanger sequencing used to confirm the presence of the targeted nitro genes. Results confirmed >95% homology for both genes.

Figure 2.7           Preparing the dsRNA reaction template for injection into C. lectularius. (a) The image of the arthropod on the left is that of C. lectularius. A genomic DNA extraction yields the template for a PCR. (b) T7 primer adapters are synthesized upstream of the intended silencing target to promote transcription of ds RNA. The first G nucleotide in the 3’ GGG codon is the first nucleotide incorporated into the dsRNA molecule. (c) a 1% agarose gel in TAE showing the raw Nitro_531_T7 PCR products used as the template for the dsRNA synthesis reaction. The intended size of the products is 531 base pairs, which is inline with lane 1 where a 1 kb ladder was run as a reference.

3.3.3 Silencing of nitro-76 and nitro-77

Out of 40 individuals injected with the combined nitro-76 and nitro-77 dsRNA injections, survival percentages at 24, 48 and 72 hr were 87.5%, 85% and 78% respectively. The injection strategy and silencing rates for 76_CiT7 and 77_CiT7 as measured with qPCR are seen in figure 2.8. Injections were repeated with dsRNA targeting the 5’ region of nitro-76 with the template designated Nitro-531-T7 (figure 2.7). Baptofect transfection reagent at 10 µM was included with Nitro_531_T7 dsRNA at 250 ng/µl and 2 µl was injected into individual bed bugs (figure 2.10). A total of 20 individuals were injected and survival for 48 and 96 hrs was 60% and 45%, respectively. At 96 hr post injection, expression levels for nitro-54, 75, 76, 77, 78 and 79 were 15.03, 0.78, 8.58e-6, 1.29e-5, 1.41 and 1.79, respectively, relative to the constitutively expressed beta-tubulin. Only the targets, nitro-76 and nitro-77 had expression levels lower than 1

Figure 2.8           Injection of preliminary nitro-76 and nitro-77 dsRNA silencing mixtures. The image on the left is that of C. lectularius with the targeted injection area outlined in dotted purple. The of focus is outlined in dotted yellow and is where the capillary needle enters the bed bug at the intersection between the abdominal paratergites. The graph on the left displays the initial silencing of nitro-76 and nitro-77 indicated by relative quantity (RQ) on the y-axis and time post injection on the x-axis. All qPCR data points were made with 4 individual bed bugs, using 2 males and 2 females.

Figure 2.9        Injection of nitro-76 and nitro-77 dsRNA with Babtofect transfection reagent. The image on the right is that of C. lectularius with the targeted injection area outlined in dotted purple. The of focus is outlined in dotted yellow and is where the capillary needle enters the bed bug at the intersection between the abdominal paratergites. The graph on the left displays the initial silencing of nitro-76 and nitro-77 indicated by relative quantity (RQ) on the y-axis and time post injection on the x-axis. All qPCR data points were made with 4 individual bed bugs, using 2 males and 2 females.

2.1.8             Primary and secondary erythema produced from individual bites

            The bite assays were done using two groups of 10 individual bed bugs injected with either dsRNA or PBS. The first group were allowed to bite the human volunteer 48 hours after injection while the second group fed 72 hours after injection. In the first group feeding times ranged from 6 min 22 sec to 25 min 5 sec with an average of 11 min 55 sec and a standard deviation of 6 min. In the second group feeding times ranged from 7 min 14 sec to 32 min 4 sec with an average of 14 min 54 sec and a standard deviation of 8 min 30 sec. The average blood meal weight was 9.54 mg for the first group with a standard deviation of 4.0 mg and the average blood meal weight for the second group was 12.37 mg with a standard deviation of 2.98 mg. The areas of erythema produced in both the nitrophorin and control bed bugs are displayed in figure 2.11 in millimeters squared and they are taken at 6, 24 and 48 hr after the initial blood meal. Additionally, the reaction erythemas were measured for males versus females, regardless of their injection status (figure 2.12).

Figure 2.10       Erythematic response of a human volunteer exposed to nitrophorin knock-down and control bed bug bites. (a) The primary erythema is the first dermal reaction that appears post blood-feeding by C. lectularius. Here, total area of the reaction is measured by ImageJ in mm2 and tracked over time. The blue line designates nitrophorin knock-down bed bugs and the red line indicated control bed bugs. (b) The secondary erythema is the darker red reaction seen at the epicenter of the bed bug bite and usually appears later. (c) Primary erythema as measured in both experimental groups fed on a human volunteer 72 hours after injections. (d) Secondary erythema as measured in both experimental groups fed on a human volunteer 72 hours after injections.

Figure 2.11       Erythematic response of a human volunteer exposed to either male or female bed bug bites. (a) The primary erythema is the first dermal reaction that appears post blood-feeding by C. lectularius. Here, total area of the reaction is measured by ImageJ in mm2 and tracked over time. The blue line designates male bed bugs and the red line indicated female bed bugs. (b) The secondary erythema is the darker red reaction seen at the epicenter of the bed bug bite and usually appears later.

Figure 2.12       Erythematic response of a human volunteer exposed bed bug bites independent of time post injection. This means that bites were administered 24 and 48 hr post injection of wither control or experimental conditions and the results were pooled. (a) The primary erythema is the first dermal reaction that appears post blood-feeding by C. lectularius. Here, total area of the reaction is measured by ImageJ in mm2 and tracked over time. The blue line designates male bed bugs and the red line indicates female bed bugs. (b) The secondary erythema is the darker red reaction seen at the epicenter of the bed bug bite and usually appears later. A total of 10 bed bugs were used for each replicate.

            The average time in minutes spent feeding by the control group 48 and 72 hours after injection with PBS was 14.53, 95% CI (±5.81) and 12.39, 95% CI (±4.14) min, respectively (figure 2.14). The average time in minutes spent feeding by the nitrophorin dsRNA-injected group 48 and 72 hours post injection was 13.85, 95% CI (±6.79) and 17.06, 95% CI (±9.62) min, respectively. When replicates from both time points are pooled, decreasing the range of our 95% CI, the average time spent feeding for the control and dsRNA injected groups was 13.46 min, 95% CI (±3.43) and 15.45 min, 95% CI (±5.65). The average amount of blood ingested in mg by the control group 48 and 72 hours after injection with PBS was 10.52, 95% CI (±3.95) and 11.94, 95% CI (±3.34) min, respectively (figure 2.14). The average amount of blood ingested in mg by the nitrophorin dsRNA-injected group 48 and 72 hours post injection was 8.56, 95% CI (±3.09) and 12.80, 95% CI (±1.95), respectively. When replicates from both time points are pooled the average amount of blood ingested for the control and dsRNA injected groups was 11.23 mg, 95% CI (±2.48) and 10.68 mg, 95% CI (±2.21).

Figure 2.13       Total ingested blood and feeding time for bed bugs either injected with dsRNA or PBS. The total feeding time in minutes is represented by the bars on the left with the control bed bugs injected with PBS seen in blue and the experimental bed bugs injected with dsRNA specific to nitrophorin on the right. The total weight of blood ingested is represented by the bars on the right with the control bed bugs injected with PBS seen in blue and the experimental bed bugs injected with dsRNA specific to nitrophorin on the right. There are a total of 10 bed bugs representing each data point and they are collected from bed bugs feeding 48 and 72 hours post injection.

Discussion

            These preliminary findings suggest that there is no significant difference between primary or secondary erythematic reactions in a single human volunteer bitten by either C. lectularius with a normal or a down-regulated nitrophorin expression level. The findings do suggest that nitrophorin dsRNA-injected bed bugs take a longer time to feed, although more evidence is needed. The time taken to feed by the nitrophorin-injected bed bugs varied greatly 72 hr after injection as indicated by the standard deviation, which was 65% of the average at 10.98 min. This could because of the variablility inherent in injections leading to variations in knock-down effectiveness. In some of the injected bed bugs, the injection mixture seeps out of the injection wound or is distributed unevenly throughout the internal anatomy. These are issues that are difficult to resolve and should be approached through more replicates and increased methodological skill in the injecting individual. If only the top 3 times are taken from the nitrophorin-injected groups, which still offers statistical significance, the time feeding is 125% and 180% higher at 48 and 72 hr post-injection, respectively, than in the control group. This is represented by an 18.13 min feeding time in the nitrophorin dsRNA-injected bed bugs at 48 hr with a 95% confidence interval of 6.21 min and a 22.29 min feeding time in the nitrophorin dsRNA-injected bed bugs at 72 hr with a 95% confidence interval of 10.31 min. In a study that achieved similar knockdown levels of nitrophorin in the triatomine bug Rhodnius prolixus (Reduviidae: Triatominae) of  >99%, they found that there was an increased time spent blood feeding in the knockdown replicates (Araujo et al., 2009). They also found that although they were able to reduce the level of haemproteins in the salivary glands by 82%, total protein levels remained the same. This suggests that there are compensatory signals triggereing the upregulation of other salivary genes to fill the role of the knocked down nitrophorins. In figure 3.10b I found that when nitro-76 and nitro-77 were knocked down, nitro-54 transcription was increased by 15-fold, suggesting a similar compensatory effect. A similar study by the same group showed that knock down of NP2 in R. prolixus at levels below 90% by RNAi increased coagulation of blood by 4-fold (Araujo et al., 2006). They also found that feeding ~13 ug of NP2-specific dsRNA to R. prolixus in the blood meal reduced gene expression levels by 42 (+/-10%). While these levels of silencing are low compared to what was achieved in the current study, they do represent a less invasive method that could potentially yield results and have not been tested in C. lectularius. One of the limitations of RNAi is that it does not directly elminate proteins, but rather mRNA tanscripts, so any proteins present in the salivary gland prior to dsRNA injection will carry out it’s role within the cell as normal. Additionally, there will be attempts by the cell to compensate for any drops in protein levels by upregulating transcription of the target gene, making precise RNAi experiments difficult. Recent technological advances overcome these challenges by completely knocking out the gene using CRISPR/Cas to engineer homozygous mutants lacking the target gene. Creating these knock-outs has become commonplace in non-model arthropods in the last 5 years (Nuss, Sharma and Gulia-Nuss, 2021), and with the creation of yolk protein-Cas9 fusion proteins, the technically demanding embryo injection is abrogated.

            These results represent the first experimental attempts to modulate the human immune response to the nitrophorins of C. lectularius. There are inherent stochastic variables involved with using a human subject that complicate the results of these data. The immunological background of the subject, defined by years of antigenic exposure, will determine the type and severity of the allergic response at the bite site. Someone preexposed to nitrophorin may react differently when bitten by a bed bug than an immunologically naïve subject. Nitrophorin was previously shown to cause a severe, hypersensitivity reaction in a single individual (Leverkus et al., 2006), which was an abberant response, as demonstated by the rarity of these hospital visits. Comprehensive epidemiological data concerning bite reactions amongst humans is hard to collect because bed bug bites often go unnoticed or ignored. Because of the copmlexity of the human immune system, future experiments testing the reactivity of nitrophorin should include an in vitro component that controls for the amounts of immune molecules in each test.

References

Araujo, R. N. et al. (2006) ‘RNA interference of the salivary gland nitrophorin 2 in the triatomine bug Rhodnius prolixus (Hemiptera: Reduviidae) by dsRNA ingestion or injection’, Insect Biochemistry and Molecular Biology. Insect Biochem Mol Biol, 36(9), pp. 683–693. doi: 10.1016/j.ibmb.2006.05.012.

Araujo, R. N. et al. (2009) ‘The role of salivary nitrophorins in the ingestion of blood by the triatomine bug Rhodnius prolixus (Reduviidae: Triatominae)’, Insect Biochemistry and Molecular Biology. Pergamon, 39(2), pp. 83–89. doi: 10.1016/j.ibmb.2008.10.002.

Balvín, O. et al. (2012) ‘Mitochondrial DNA and morphology show independent evolutionary histories of bedbug Cimex lectularius (Heteroptera: Cimicidae) on bats and humans’, Parasitology Research. Parasitol Res, 111(1), pp. 457–469. doi: 10.1007/s00436-012-2862-5.

Booth, W. et al. (2015) ‘Host association drives genetic divergence in the bed bug, Cimex lectularius’, Molecular Ecology. Blackwell Publishing Ltd, 24(5), pp. 980–992. doi: 10.1111/mec.13086.

Doggett, S. L. et al. (2012) ‘Bed bugs: clinical relevance and control options’, Clinical Microbiology Reviews. Clin Microbiol Rev, 25(1), pp. 164–192. doi: 10.1128/CMR.05015-11.

Francischetti, I. M. B. et al. (2010) ‘Insight into the sialome of the bed bug, Cimex lectularius.’, Journal of Proteome Research. NIH Public Access, 9(8), pp. 3820–31. doi: 10.1021/pr1000169.

Goddard, J. and Edwards, K. T. (2013) ‘Effects of bed bug saliva on human skin’, JAMA Dermatology. American Medical Association, 149(3), p. 372. doi: 10.1001/jamadermatol.2013.878.

Goddard, J., Edwards, K. T. and de Shazo, R. D. (2011) ‘Observations on development of cutaneous lesions from bites by the common bed bug, Cimex lectularius L. ’, Midsouth Entomologist, 4(2), pp. 49–52.

Kõressaar, T. et al. (2018) ‘Primer3-masker: integrating masking of template sequence with primer design software’, Bioinformatics. Oxford University Press, 34(11), pp. 1937–1938. doi: 10.1093/bioinformatics/bty036.

Leverkus, M. et al. (2006) ‘Bullous allergic hypersensitivity to bed bug bites mediated by IgE against salivary nitrophorin’, Journal of Investigative Dermatology, 126(1), pp. 91–96. doi: 10.1038/sj.jid.5700012.

Omori, N. (1941) ‘Comparative studies on the ecology and physiology of common and tropical bed bugs, with special reference to the reactions to temperature and moisture.’, Journal of the Medical Association of Formosa, 40(3), pp. 555–636.

Potter, M. F. (2004) ‘Bedbugs: understanding and control’, Pest Control Technology, 32(8), pp. 13–17.

Potter, M. F. (2006) ‘The perfect storm: an extension view on bed bugs’, American Entomologist, 52(2), pp. 102–104.

Reinhardt, K. et al. (2009) ‘Sensitivity to bites by the bedbug, Cimex lectularius’, Medical and Veterinary Entomology. Med Vet Entomol, 23(2), pp. 163–166. doi: 10.1111/j.1365-2915.2008.00793.x.

Reinhardt, K., Isaac, D. and Naylor, R. (2010) ‘Estimating the feeding rate of the bedbug Cimex lectularius in an infested room: An inexpensive method and a case study’, Medical and Veterinary Entomology. Med Vet Entomol, 24(1), pp. 46–54. doi: 10.1111/j.1365-2915.2009.00847.x.

Schwartz, L. B. (2004) ‘Effector cells of anaphylaxis: mast cells and basophils ’, Novartis Foundation Symposium, 257, pp. 65–74.

Sheele, J. M. et al. (2017) ‘Antibody and cytokine levels in humans fed on by the common bedbug, Cimex lectularius L’, Parasite Immunology, 39(3), p. e12411. doi: 10.1111/pim.12411.

Sheele, J. M. et al. (2019) ‘Cimicosis in persons previously fed upon by bed bugs’, Cureus. Cureus, Inc., 11(10). doi: 10.7759/cureus.5941.

Sheele, J. M. et al. (2020) ‘Human immunoglobulin G responses to Cimex lectularius L. saliva’, Parasite Immunology. Blackwell Publishing Ltd, 42(12), p. e12764. doi: 10.1111/pim.12764.

Sutton, B. et al. (2019) ‘IgE antibodies: from structure to function and clinical translation’, Antibodies. MDPI AG, 8(1), p. 19. doi: 10.3390/antib8010019.

CHAPTER III 

GENERATING TRANSGENIC KNOCKOUT LINES OF AEDES ALBOPICTUS FOR THE FUNCTIONAL CHARACTERIZATION OF OR4 

Background

            Aedes albopictus is arguably the major mosquito threat facing the American public (Dieme, Ciota and Kramer, 2020) and is a major threat throughout most of the world. This was highlighted by concerns during the ongoing Zika virus epidemic (McKenzie, Wilson and Zohdy, 2019). While a related species, Ae. aegypti, is a more capable vector for many diseases due to its strong preference for feeding on humans, Ae. albopictus has displaced Ae. aegypti throughout most of the continental U.S. over the last three decades and is a capable disease vector (Hopperstad, Sallam and Reiskind, 2021). Similar situations of range expansion by Ae. albopictus have occurred throughout the world. Several factors have led to Ae. albopictus’ success as an invasive species such as their abilities to outcompete other Aedes species (Liu, Tian and Ruan, 2020), blood-feed on a variety of hosts, and adapt to cooler climatic conditions. These factors likely involve active niche shifting (Cunze et al., 2018) and it is hypothesized that genetic manipulation of factors related to Ae. albopictus success could eventually lead to the production of a strain of mosquito that would be less dangerous to people, or to knowledge that could be incorporated into conventional insect control strategies. The newly available CRISPR/Cas9 genome editing strategies, coupled with established phenotypic assays are employed to work towards testing this hypothesis. First, this species offers several clear genetic targets involved in niche-preference that have been recently uncovered and remain largely unstudied. Second, Ae. albopictus is a major threat to the health of the American public and CRISPR/Cas9-based genome engineering has not been reported for this species to date. Third, Ae. albopictus’ presence in the continental U.S. likely keeps the most capable human disease vector, Ae. aegypti, in check. So, it could be argued that Ae. albopictus is not a good target for elimination, but rather that niche-preference engineering could be used to change it into a less competent vector while allowing it to continue to repress a more dangerous vector.

            Host-seeking behavior is an important component of Ae. albopictus’s ability to invade diverse ecological systems and this behavior is largely driven by their chemosensory system (Yan et al., 2020). In insects, odors are perceived as volatile molecules binding to receptors exposed to the extracellular surface of chemosensory neurons, which include odorant receptors (ORs), ionotropic receptors (IRs), gustatory receptors (GRs), and other, more unique, receptors such as the Pickpocket (PPK) and TRP receptors (Joseph and Carlson, 2015). There is often only one odorant receptor expressed per chemosensory neuron, which facilitates the recognition of a wide variety of odors. Identical neurons project their axons to the same glomeruli within the antennal lobe where they will synapse with higher-order neurons (Komiyama and Luo, 2006). In Ae. aegypti, the evolution of anthropophilic behavior has been linked to the odorant receptor, AeOR4 (McBride et al., 2014). The experiments outlined here detail the targeting of the AeOR4 ortholog in Ae. albopictus for homozygous knockout by the Cas9 endonuclease.

Material and methods

3.1.1             Mosquito colony maintenance

            Aedes albopictus Foshan strain were obtained from MA4 in a square petri dish containing DI water-soaked filter paper. Eggs were floated in DI water at 27°C in a plastic container and placed in a vacuum chamber. Careful care was taken to ensure that all eggs were submerged in the water to maximize hatch rates. Air was removed from the vacuum chamber by the application of an air pump for 5 mins and the container was left in the vacuum chamber for 6 hours. The container may need more time in the vacuum chamber to completely de-oxygenate the water if the eggs do not hatch. Over 24 hours in the vacuum chamber was never exceeded to prevent the death of the mosquito larvae.

3.1.2             sgRNA synthesis

            Single guide RNAs were selected based on their homology to the genetic background of colony mosquitos reared in our insectary. The Or4 gene (AaeL_AAEL015147) homolog previously characterized in Ae. aegypti was used as an input for a BLAST search of the non-redundant database with Ae. albopictus designated as the “Organism” in the “Choose search set” dropdown menu. Primers were selected for both genes flanking the 5-prime untranslated region of the gene and extending at least 2,000 base pairs 3-prime to include most of the coding exons. Primer3 was used to design the primers.

3.1.3             Construction of the pressurized injection system

            A 13.3 kg, 7.8 L pressurized tank containing nitrogen gas was used as the air source to drive flow through the capillary system. The regulator on the tank was a TriTech microINJECTOR™ with an inlet and outlet pressure gauge and a 1000 PSI maximum outlet capacity. Connections to all the components within the system were made using Clippard 0.25 OD, 0.125 ID polyurethane tubing (URH1-0804). The air supply first enters the piston driven World Precision Instruments (WPI) PV830 Pneumatic PicoPump through the inlet at the back of the instrument. Air entering the system never exceeded 200 PSI to prevent blowing out the pump. The eject port of the pump was connected to a WPI capillary tube holder (MPH3-10) using polyurethane tubing. The tube holder was housed on a custom-built threaded brass rod to facilitate movement of the connected needle between the injection and beveling stations without needing to move the entire system.

Figure 3.1           Injection setup with an emphasis on the path of the gas from the tank to the capillary needle holder. (a) The Tri-tech regulator controls the pressure of the gas coming from the tank. A slight turn clockwise will set the pressure to 40 psi, which is sufficient for injection. (b) The back panel display of the PV830 Pneumatic PicoPump showing the input tubing coming from the tank regulator. As noted, the PSI should never exceed 150 PSI to avoid blowing out the pump. (c) The final outlet of the gas is at the microcapillary holder, which is shown here without the capillary needle. (d) The WPI PV830 Pneumatic PicoPump controls the amount of gas ejected at the capillary with a pressurized solenoid. The tubing is secured to the ejection port here with parafilm. All junctions are circled with a dotted red line.

3.1.4             Construction and cloning of knock-in plasmid

            Genomic DNA from a single colony mosquito was collected to reduce heterogeneity in the target gene. Pigments in the mosquito eye may inhibit downstream PCR reactions so the head of all mosquitos should be removed prior to DNA extraction. The Monarch® Genomic DNA Purification Kit (T3010S) was used for genomic DNA extraction. Briefly, a single, fresh mosquito was suspended in 200 µl of modified Tissue Lysis Buffer and 10 µl of Proteinase K in a 1.5 ml microcentrifuge tube and ground with a sterilized plastic pestle. The digestion was incubated at 56°C and 1400 RPMs for 3 hrs. The extended incubation time ensures that all residual RNA has been degraded and that the optimum cell lysis has been achieved. The digest was centrifuged at room temperature and 12,000 gs for 3 mins to pellet the cellular debris and the supernatant was transferred to a new 1.5 ml microcentrifuge tube. The supernatant was combined with 3 µl of RNase A, vortexed and incubated with 1,400 RPM agitation at 56°C for 5 mins to completely remove all residual RNA from the lysate. Add 400 µl of gDNA Binding Buffer to the lysate and vortex at maximum speed for 10 seconds. The lysate/buffer mix was then transferred to gDNA purification column and centrifuged for 3 mins at 1,000 g to bind the gDNA to the column. This was followed by centrifugation at 12,000 g for 1 min to clear the column of the binding buffer/lysate mix. The flowthrough was discarded, and 500 µl of gDNA wash buffer was added to the column and centrifuged for 1 min at 12,000 g. The flowthrough was discarded and 50 µl of RNase/DNase free water pre-heated to 65°C was added directly to the column and incubated for 5 mins at room temperature. The column was placed in a sterile 1.5 ml microcentrifuge tube and centrifuged at 12,000 g for 1 min. The resulting purified genomic DNA was the quantified using the Nanodrop-C Spectrophotometer.

Figure 3.2           Depiction of the Gibson Assembly reaction. (1) Primers are designed with a specificity to the target and the designated annealed fragment. (2) The Gibson Assembly begins with a T5 exonuclease reaction which de-polymerizes the double-stranded DNA duplexes starting at the 5’ end. (3) The continuation of the 5’ exonuclease activity. (4) DNA polymerase comes into seal newly annealed fragments where the T5 did not excise the nucleotides. (5) Ligase comes in to bind. (6) The final assembled plasmid.

            HA and Gibson assembly plasmids were cloned together in two separate NEB® 10-beta Competent E. coli (High Efficiency) reactions (C3019H) using the high efficiency transformation protocol. For each, 50 µls of 10-beta competent cells stored at -80°C were thawed, on ice, for 10 mins until the last of the ice disappears. 2 µl from each reaction was added to the thawed cells and the tubes were carefully flicked 4 times with no use of the vortex mixer. The mixtures were then incubated on ice for 30 mins with no mixing. Cells were then heat shocked at 42°C for exactly 30 secs in a thermocycler and moved to ice for an additional 5 min incubation. This was combined with 950 µl of NEB 10-beta/Stable Outgrowth Medium at room temperature and incubated at 37°C for 60 mins at 250 RPMs. The transformed cells were diluted 10-fold in the outgrowth media and both dilutions were plated on pre-warmed LB agar selection plates containing 0.1 mg/ml ampicillin, 0.2 mg/ml X-gal and 1 mM IPTG. For each transformation, 50 µls was pipetted onto the selection plates using wide-bore pipette tips, which was spread using a sterilized plastic media spatula and incubated overnight at 37°C. Transformation efficiencies were calculated using the following equation:

Equation 1 Calculation used to determine the efficiency of plasmid transformation by the desired organism

White colonies were picked using a sterile 10 µl pipette tip which was dipped in 200 µl PCR tubes containing 5 µl of sterile water for colony PCR before being placed in 5 mls of liquid LB broth containing 0.1 mg/ml ampicillin in 15 ml Falcon tubes. Colony PCR was performed using the REDpSeq1 and HAp19Seq primers for the pEF1dsRED and pHA plasmids, respectively. Q5® High-Fidelity DNA Polymerase (M0491) was used for PCR of both plasmids being screened for assembly. Q5 reactions were assembled on ice and contained 200 uM dNTPs, 0.5 uM forward and reverse primers, 100 ng plasmid template and 0.02 U/µl of Q5 polymerase. The pEF1dsRED template was incubated in the thermocycler under the following conditions: 1 cycle at 98°C for 30 secs, 35 cycles of 98°C for 10 secs, 64°C for 15 secs and 72°C for 70 secs and 1 cycle of 72°C for 2 mins. The pHA template was incubated in the thermocycler under the following conditions: 1 cycle at 98°C for 30 secs, 35 cycles of 98°C for 15 secs, 66°C for 15 secs and 72°C for 40 secs and 1 cycle of 72°C for 2 mins. Both PCR reactions were run on a 1% agarose gel at 90 V for 30 mins and screened for the presence of a 1,019 and 718 base pair band for pEF1dsRED and pHA, respectively. Positive colony cultures were used for further processing and plasmid purification.

3.1.5             Injection of embryos

            The first step in embryo injections is to collect viable embryos from the target species. Adult mosquitos were fed blood from a membrane feeder wrapped with a tight film of parafilm and filled with blood, which was heated by circulating water at 37°C. The colony was starved for 48 hr to kill all the unfed females and males. Three days after feeding, eggs were collected from female mosquitos. A 50 ml conical tube was used to collect females who were placed in the dark at 27°C for 25 min with a filter paper submerged in water used as an oviposition pad. The females were removed from the tube and separated to a new colony. The eggs were collected from the filter paper and transferred to a new piece of moist filter paper aligned against a microscope cover slip. From the point the eggs were separated from the females, only 20 min passes prior to affixing the eggs to the cover slip for injection. Once the eggs are aligned against the cover slip, with the proximal, pointed end facing out, all of the water from the filter paper is sopped up with a dry piece of filter paper. It is essential that the filter paper is completely dried prior to removing the cover slip to ensure the eggs remain in a straight line. The cover slip was pulled away from the eggs and another cover slip with a piece of attached Tegaderm is lightly pressed against the eggs to affix them. They are covered in 50 ul of Halocarbon-50 and prepared for injection.

            Injections were carried out under a stereomicroscope manually affixed to an adjustable stage using epoxy resin. The injection capillary needle was filled immediately prior to injection using a Microcap tube manually pulled over the flame of a Bunsen burner. The Microcap tube will pull up the injection mix through capillary action and only needs less than 1 ul of mix to inject up to 50 embryos. The loaded capillary needle was connected to the pressurized injection setup and the tip was carefully set under the microscope pointed toward the proximal tip of the embryos, submerged in Halocarbon-50. The PicoPump is set to 20 kPa and a test injection determined if the ejected volume was sufficient as to not over or underfill the embryo. Once the desired pressure is set, the eggs are injected one by one, making sure that any eggs not injected properly are removed to avoid laborious screening procedures after hatching. The Halocarbon-50 was removed from the eggs using DI-water and they were allowed to develop at 27°C in a square petri dish at 85% relative humidity for 48 hr after which, standard hatching procedures were carried out.

Results

3.1.6             Construction of plasmids in silico

            A BLAST search of the non-redundant nucleotide database gave 2 probable hits in Ae. albopictus. LOC109400239, Aedes albopictus odorant receptor 4-like, had a total score of 837 on a query coverage of 76% and a percent identity of 79.27% with an E value of 0.0. LOC109423565, Aedes albopictus odorant receptor 4-like, had a total score of 832 on a query coverage of 74% and a percent identity of 79.51% with an E value of 0.0. Primers designed to

Figure 3.3           A diagrammatic representation of the two OR4 genes in Ae. albopictus. (a) LOC109400239 shows the 4 exons contained in the 2,030 bp coding sequence in blue. The yellow box represents the zoomed in region on the bottom of the figure and is the region targeted by the CRISPR/Cas target using the sgRNA, which is specific to a 20 base pair region in the coding sequence. (b) LOC109423565 shows the 6 exons in blue. Note the 56,883 base pair intronic region between exon 5 and 6. The Diagram at the bottom displays the 4 regions targeted for sequencing with each yellow bar representing one set of primers.

specifically amplify LOC109400239. Sanger sequencing and MEGA alignment of the 4 sets of OR4 homology arm PCR templates revealed that the OR4 gene in our Ae. albopictus colony has an alignment to LOC109423565 with an E-score of 0.0, a max score of 1,282, a total score of 2290 and an identity of 90.51%.  It also aligns to LOC109400239 with an E-score of 0.0, a max score of 687, a total score of 1639 and an identity of 94.75%.

3.1.7             Synthesis and assembly of component DNA fragments and plasmids

Figure 3.4           Gibson assembly producing the pEFdsRED plasmid. (a) Qiagen miniprep plasmid products run from 3 separate colonies of beta-10 E. coli chemically transformed with a 4 fragment Gibson assembly. The bands show the characteristic 3 band profile of a circular plasmid with the supercoil running at ~1.5 kb. (b) A diagrammatic representation of the pEFdsRED plasmid once assembled. The dsRED marker is the coding sequence flanked by a 5’EF1alpha promoter and 3’EF1alpha terminator, both endogenous to Ae. albopictus. The plasmid is designed with restriction sites in the F_5’EF1a and R_3’EF1a primers, flanked by the Gibson overhang adapters, to facilitate a subsequent cloning reaction.

            The Gibson Assembly joined 0.2 pmol of each of the 4 fragments: 5’ EF1a, dsRED, 3’ EF1a and pUC19 backbone. Five transformed, white colonies were picked for miniprep and yielded between 120 and 368 ng/µl for each. The plasmids run aon a gel were screened and the consistent bands that showed positive for the sequencing junction were selected for digestion, clean up, ligation, clean up and transformation. The final pEFaOR4dsRED plasmid is seen in 3.4 and the gel of 5 of the miniprepped samples were ran on a gel, seen in figure 3.4, b. The final plasmid is used as the injection template for Ae. albopictus.

Figure 3.5           Restriction digest, ligation and cloning producing the pHA plasmid. (a) Qiagen miniprep plasmid products run from 3 separate colonies of beta-10 E. coli chemically transformed with a 2-fragment ligation. The bands show the characteristic 3 band profile of a circular plasmid with the supercoil running at ~1.2 kb. (b) A diagrammatic representation of the pHA plasmid once assembled. The HA fragment is homologous to the target gene, OR4, and contains the gRNA site roughly in the middle of its sequence. The cargo to be inserted into the genome of Ae. albopictus is inserted between this opened plasmid in a subsequent cloning reaction.

Figure 3.6           The final injection plasmid targeting the OR4 gene in Ae. albopictus. (a) pEFaOR4dsRED is 8,205 base pairs long and contains a dsRED marker gene flanked by the EF1a 5’ and 3’ untranslated regions to drive constitutive expression in Ae. albopictus. The marker construct is flanked by two 1,000 base pair homology a°rms homologous to odorant receptor-4 in Ae. albopictus and driving homology-directed repair to knock-in the dsRED marker gene. (b) The final plasmid is shown in lanes 2-4 with the characteristic 3 band pattern indicative of an uncut plasmid.

3.1.8             Preliminary injections

            Of the 332 eggs injected and brought to the hatching chamber, 0% survived. It is important to note that not all eggs laid by females were injected and that the experiments were not limited by oviposition. Rather, limitations were in the time allowed for lining up and injecting the eggs within the 20 min allotted. There was an average of 47 eggs lined up during each injection procedure through the course of 7 injection runs. These results were unexpected and there needs to be methodological adjustments to increase hatch rates.

Discussion

            Preliminary injections suggest that some component within the injection mix may be lethal to the developing embryo, or that knocking out AeaeOR4 may be lethal in and of itself. However, a very important caveat is that more injections are needed to conclude this, as there was insufficient time to perform control embryo injections as that was not an experimental objective prior to performing this research. While undergoing training to perform these techniques, it was assumed that injection methodology would not lead to lethality in the embryos. As these techniques are being implemented for the first time in this lab and by this group of personnel, it is suggested and indeed necessary to perform control injections containing reaction buffer alone to validate technique and equipment. Through the process of developing these techniques, several novel methodologies were developed and fine-tuned, which does contribute to the field of embryo injection.

            In Ae. aegypti, AaegOR4 is involved with anthropophilic behavior during blood feeding (McBride et al., 2014). A BLAST search using AaegOR4 as the query identified two very similar homologs in Ae. albopictus, LOC109423565 and LOC109400239. By sequencing the targeted gene, I found that Or4 in our colony Foshan strain aligned to LOC109423565 with a 91% identity and a score of 1420 while it aligned to LOC109400239 with an 81% identity and a score of 715. Using this, LOC109400239 was designated as the homolog to target for functional analysis through mutation. There were multiple potential sgRNA target sites identified by CHOPCHOP that were centrally located in LOC109423565 for the introduction of double-stranded breaks by Cas9 (Labun et al., 2019). The machine learning driven CRISTA scoring algorithm (Abadi et al., 2017) I selected the sgRNA with the highest score from the CHOPCHOP candidates, designated here OR4.1sgRNA. Additionally, the target was chosen based on the lack of heterozygosity at the target based on single peak chromatograms. There is no ambiguity between the 26-base pair sequence in our Sanger sequencing results and the selected sgRNA target site, preventing mutations from disrupting specificity. With this information, future injections can proceed using the same sgRNA as everything constructed downstream is dependent upon it and any change would mean changing the entire knock-in plasmid.

            When constructing novel plasmids, there are several hurdles to achieving rapid and accurate assembly of the desired product. First, there is little room for leeway when attaching a promoter region to a coding sequence. The promoter must be in-frame with the ATG start codon and the primer must contain the exact sequence of the 3’-end of the promoter and the 5’-start of the coding sequence for the Gibson assembly to work properly. This was an issue when amplifying the 5’EF1-alpha promoter region from Ae. albopictus genomic DNA. Multiple polymerase chemistries were attempted including OneTaq, Phusion and Q5, all with primer annealing temperature gradients and varying concentrations of buffer enhancer and MgCl2 concentrations. Non produced the desired 2,323 bp fragments required for Gibson assembly. The fragment is required at 2 pmol concentrations for a 4-fragment Gibson assembly, so well over 100 ng was necessary, which is troubling considering the reaction did not work at all. The solution to this was to order the sequence from an outside commercial company (Gene Universal) that offered in vitro synthesis of the fragment for under 500 dollars. This would have been prohibitively expensive 10 years ago, so this is one example of where contemporary commercial offerings provided a simple solution to the problem. Another issue is codon optimization. The coding sequence for the trans-marker gene should have a codon repertoire that matches the ideal composition of tRNAs in the particular species. In this case, there was no codon optimized dsRED for Ae. albopictus but there was one that had proven to work in Ae. aegypti, a relative of our species. Whether they were close enough relatives to produce enough expression to produce an observable phenotype is yet to be observed and needs to be noted moving forward. A solution to this would be to use qPCR primers to observe undetectable levels of transcription that don’t necessarily need to proceed to translation, overcoming the need to use any tRNAs.

            Considerations during physical injection include proper climatic conditions in the injection room and post-injection rearing conditions. All the techniques used here were learned during a training course at the University of Maryland and in association with the Insect Transformation Facility. Many of those techniques are outlined in a recently released protocol (Meuti and Harrell, 2020), however this publication is intended for use in Culex spp, which requires very specific conditions during several key steps. For example, Ae. albopictus eggs are not deposited on floating egg rafts and need to be handled differently when lining up immediately prior to injection. Standard operating procedures during these experiments need to include the recording of time, temperature, and humidity so that survival rates can be better correlated to specific conditions.

References

Abadi, S. et al. (2017) ‘A machine learning approach for predicting CRISPR-Cas9 cleavage efficiencies and patterns underlying its mechanism of action’, PLoS Computational Biology. Public Library of Science, 13(10), p. e1005807. doi: 10.1371/journal.pcbi.1005807.

Cunze, S. et al. (2018) ‘Niche conservatism of Aedes albopictus and Aedes aegypti - Two mosquito species with different invasion histories’, Scientific Reports. Nature Publishing Group, 8(1), p. 7733. doi: 10.1038/s41598-018-26092-2.

Dieme, C., Ciota, A. T. and Kramer, L. D. (2020) ‘Transmission potential of Mayaro virus by Aedes albopictus, and Anopheles quadrimaculatus from the USA’, Parasites and Vectors. BioMed Central Ltd, 13(1), pp. 1–6. doi: 10.1186/s13071-020-04478-4.

Hopperstad, K. A., Sallam, M. F. and Reiskind, M. H. (2021) ‘Estimations of fine-scale species distributions of aedes aegypti and aedes albopictus (Diptera: Culicidae) in Eastern Florida’, Journal of Medical Entomology. Oxford University Press, 58(2), pp. 699–707. doi: 10.1093/jme/tjaa216.

Joseph, R. and Carlson, J. (2015) ‘Drosophila chemoreceptors: A molecular interface between the chemical world and the brain’, Trends in genetics : TIG. Trends Genet, 31(12), pp. 683–695. doi: 10.1016/J.TIG.2015.09.005.

Komiyama, T. and Luo, L. (2006) ‘Development of wiring specificity in the olfactory system’, Current opinion in neurobiology. Curr Opin Neurobiol, 16(1), pp. 67–73. doi: 10.1016/J.CONB.2005.12.002.

Labun, K. et al. (2019) ‘CHOPCHOP v3: Expanding the CRISPR web toolbox beyond genome editing’, Nucleic Acids Research. Oxford University Press, 47(W1), pp. W171–W174. doi: 10.1093/nar/gkz365.

Liu, Z., Tian, C. and Ruan, S. (2020) ‘On a network model of two competitors with applications to the invasion and competition of aedes albopictus and aedes aegypti mosquitoes in the United States’, SIAM Journal on Applied Mathematics. Society for Industrial and Applied Mathematics Publications, 80(2), pp. 929–950. doi: 10.1137/19M1257950.

McBride, C. S. et al. (2014) ‘Evolution of mosquito preference for humans linked to an odorant receptor.’, Nature. NIH Public Access, 515(7526), pp. 222–7. doi: 10.1038/nature13964.

McKenzie, B. A., Wilson, A. E. and Zohdy, S. (2019) ‘Aedes albopictus is a competent vector of Zika virus: A meta-analysis’, PLoS ONE. Public Library of Science, 14(5), p. e0216794. doi: 10.1371/journal.pone.0216794.

Meuti, M. E. and Harrell, R. (2020) ‘Preparing and injecting embryos of culex mosquitoes to generate null mutations using crispr/cas9’, Journal of Visualized Experiments. Journal of Visualized Experiments, 2020(163), pp. 1–19. doi: 10.3791/61651.

Yan, H. et al. (2020) ‘Evolution, developmental expression and function of odorant receptors in insects’, The Journal of experimental biology. J Exp Biol, 223(Pt Suppl 1). doi: 10.1242/JEB.208215.

CHAPTER IV 

CONCLUSION

            This dissertation expands on our current understanding of the sialome in C. lectularius, demonstrates knock-down of salivary nitrophorin by levels  >99.99% expands on methodologies used for embryo injections in Ae. albopictus. A comprehensive, next-generation Illumina RNA-seq library of differentially expressed salivay transcripts in C. lectularius identified 19,269 unique transcripts from the 697.9 Mb available genome. With a total of 64,721,981 paired end, 50 base pair reads, which represents 10x sequencing coverage. From these differentially expressed genes, the profile of salivary nitrphorins was definitively described as a clade of 3 groups of 2 related genes, for a total of 6 unique nitrophorin genes. All of them have a signal peptide and the log2 fold-change ranges from 13.46 to 16.40 for inositol polyphosphate 5-phosphatase K-like (LOC106662978) and 72 kDa inositol polyphosphate 5-phosphatase-like (LOC106662976), respectively, when copmared to transcripts expressed in the carcass of C. lectularius. It was previously believed that there were 8 unique salivary nitrophorins, although this was based on sequence redundancy produced from partial expressed sequence tags. With this information, future studies can focus on this set of 6 salivary nitrophorins in C. lectularius.

            In this study, two nitrophorins were found that were phylogenetically separated based on their Cys-60 and annotated as 72 kDa inositol polyphosphate 5-phosphatase-like (LOC106662976) and 72 kDa inositol polyphosphate 5-phosphatase-like (LOC106662977) had an at least 8-fold higher expression profile than the other 4 nitrophorins. These two nitrophorins reached their peak levels in newly emerged adult bed bugs that were starved for 14 days. Using RNAi, it was poossible to silence these nitrophorins by targeting the 5’ region of the genes with dsRNA. Silencing was achieved at levels of over 99.99% at 48 and 96 hours after injection with dsRNA. This represents the first time a salivary nitrophorin was silenced to these levels in C. lectularius and paves the way for the silencing of other salivary genes in this species moving forward. Another potential for silencing is nudix hydrolase (LOC106666860), which has a log2 fold expression upregulation of 20.08, making it one of the highest expressed genes in the salivary gland. This nudix hydrolase has never before been implicated in elliciting hypersensitivity reactions but it is most likely involved in anti-coagulation and may impact feeding behavior when silenced. Silencing the two Cys-60 nitrophorins proved to have little to no impact on blood feeding behavior or in their ability to differentially ellicit erythema in a human volunteer. There were some outliers that had increased feeding times, suggesting that more biological replicates might yield some more consistent and significant results. Future studies should focus on silencing all 6 salivary nitrophorins and examing the impact on feeding behavior. Additionally, more needs to be done to look at the levels of protein in silenced bed bugs,a s trancriptional silencing may not mean a depletion in protein levels.

            The majority of work done in this report towards silencing of OR4 in Ae. albopictus builds on methodologies used for embryo injections. Previous experiments using PiggyBac-based transgenesis achieved mutation efficiencies of 1-2%. From the 332 eggs injected here, non of them were viable. Observations suggest that successful injections are reliant on the quality of the borosilicate capillary needle used for injection. Any deviations from perfect beveling will lead to embryonic fluid seeping from the eggs after injections, ensuring the death of the egg. Future experiments should initially focus on producing knock-out mutants through non-homologous end joining first, before attempting an HDR-driven konck-in. This will ensure that the sgRNA is efficient at targeting and that the knock-out does not produce a lethal phenotype. An injection mix including dsRNA targeting ku70 and a knock-in plasmid has been developed but this is required to be tested by more injections to achieve success or definitively say that the gene is essential for embryo viability.

            Overall, the methodologies demostrated here represent how current molecular techniques can contribute to our understanding of arthropods genetics. Illumina currently offers the greatest depth of short-read sequencing technologies and produced here a 19,269 salivary transcriptome in C. lectularius. Using RNAi, it was possible to silence the highest expressed salivary nitophorins by >99.99%. This sets a precedent for further RNAi-based knock-down experiments in the salivary glands of C. lectularius. Additionally, a CRISPR/Cas-based knock-in template was rapidly and affordably assembled facilitating future targeted mutagenesis experiments in Ae. albopictus.

[1] Bayesian phylogeny (5M generations) of all Anopheles gambiae genes and all other known salivary C-type lectins from mosquitoes, all Rhodnius, Triatoma, other related heteropterans and a broad range of metazoans from a previous phylogenetic construction of all known lysozymes (Callewaert and Michiels 2010). Two protein sequences were removed due to interference with the alignment process (Ixodes_XP_00243606 and AGAP005717). Weak backbone support within C-type lysozymes. “Arthropods” refers to groups with mixed subclasses of arthropods. Cimex I-type lysozyme is placed within the insecta. B) Partial MA plot showing the high-salivary gland expression levels of five related Cimex lysozymes. C) Alignment of all protein sequences from tree in figure 4A (about 10% of variable N-terminus removed for display purposes).  D) Alignment of the boxed region from figure 4C for all Cimex genes and the immunity related lysozymes 3 and 4 from Harmonia. The two Cimex genes with predicted muramidase activity are outlined in the blue box. The catalytic glutamic acid (E) and aspartic acid (D) residues necessary for muramidase activity are shown in blue text.

[2] These genes do not contain the Cys-60 residue believed to be integral to heme-binding, however they are predicted to have an otherwise suitable heme-binding pocket. (A) Bayesian consensus phylogeny (MrBayes 5M generations using amino acid sequences) of all related published and unpublished Heteropteran salivary proteins, a selection of related IPP5P-like proteins from invertebrates, and an outgroup comprised of the two major groups of IPP5P proteins present in metazoans. The clade labeled “Saliva/Venom” are IPP5P-related proteins found in the saliva or venom of various Heteroptera. All of these genes contain potential cleavage sites, which implies they may be signal peptides, as reported by SignalP. Cimex species other than C. lectularius are derived from partial sequences. Hemipteran data labelled “unpublished” were supplied by King. The two major, strongly supported, clades of genes with IPP5P function are named “E” and “B” following the vertebrate nomenclature. Synapotojanin proteins contain a region of high homology to IPP5Ps in insects but were not included as their overall structure includes several other unrelated modular domains. (B) All IPP5P-like genes significantly enriched in the salivary glands versus whole body minus salivary glands. (C) Homology modelling using the Swiss-Model Workspace published CleNitro1 template on top and an overlay of all 6 proposed nitrophorins with the associated heme ligand in the binding pocket below.