Optimized genetic tools for neuroanatomical and functional mapping of the Aedes aegypti olfactory system

Abstract

The mosquito Aedes aegypti is an emerging model insect for invertebrate neurobiology. We detail the application of a dual transgenesis marker system that reports the nature of transgene integration with circular donor template for CRISPR–Cas9-mediated homology-directed repair at target mosquito chemoreceptor genes. Employing this approach, we demonstrate the establishment of cell-type-specific T2A-QF2 driver lines for the A. aegypti olfactory co-receptor genes Ir8a and orco via canonical homology-directed repair and the CO₂ receptor complex gene Gr1 via noncanonical homology-directed repair involving duplication of the intended T2A-QF2 integration cassette separated by intervening donor plasmid sequence. Using Gr1+ olfactory sensory neurons as an example, we show that introgression of such T2A-QF2 driver and QUAS responder transgenes into a yellow cuticular pigmentation mutant strain facilitates transcuticular calcium imaging of CO₂-evoked neural activity on the maxillary palps with enhanced sensitivity relative to wild-type mosquitoes enveloped by dark melanized cuticle. We further apply Cre-loxP excision to derive marker-free T2A-QF2 in-frame fusions to clearly map axonal projection patterns from olfactory sensory neurons expressing these 3 chemoreceptors into the A. aegypti antennal lobe devoid of background interference from 3xP3-based fluorescent transgenesis markers. The marker-free Gr1 T2A-QF2 driver facilitates clear recording of CO₂-evoked responses in this central brain region using the genetically encoded calcium indicators GCaMP6s and CaMPARI2. Systematic application of these optimized methods to different chemoreceptors stands to enable mapping A. aegypti olfactory circuits at peripheral and central levels of olfactory coding at high resolution.

mosquito, CRISPR–Cas9, olfaction, chemoreceptor, neurogenetics

Introduction

The yellow fever mosquito Aedes aegypti is a globally important vector of yellow fever, dengue, Zika, and chikungunya viruses. These mosquitoes use their highly tuned sense of smell to detect humans, find sources of nectar, and locate oviposition sites. Given these complex olfactory behaviors, including epidemiologically relevant ones such as anthropophilic host-seeking behavior, A. aegypti has become an important insect model to study chemosensation and sensory integration (DeGennaro et al. 2013; McBride et al. 2014; McMeniman et al. 2014; van Breugel et al. 2015; Duvall et al. 2019; Raji et al. 2019; Jové et al. 2020; Melo et al. 2020; De Obaldia et al. 2022; Herre et al. 2022; San Alberto et al. 2022; Chandel et al. 2024; Rouyar et al. 2024). Studies of A. aegypti sensory neurobiology have been catalyzed directly as the result of a high-quality genome annotation (Matthews et al. 2018), comprehensive neurotranscriptomic datasets (Matthews et al. 2016; Herre et al. 2022), and the application of genome engineering tools including the QF2/QUAS binary expression system and genome editing tools such as the CRISPR–Cas9 system to assess gene function in this mosquito species (Kistler et al. 2015; Li et al. 2017; Matthews et al. 2019; Herre et al. 2022).

The olfactory system of A. aegypti consists of 3 major olfactory appendages including the antennae, the maxillary palps, and the labella of the proboscis. Lining these organs are various morphological classes of porous sensilla (McIver 1972, 1973, 1978) that house the dendritic processes of typically 2–3 olfactory sensory neurons (OSNs) that detect diverse structural classes of volatile odorants. Large chemoreceptor gene families implicated in detection of various components of human scent and other ethologically relevant odorants are encoded in the A. aegypti genome (Matthews et al. 2018) and are expressed by olfactory appendages on the head (Herre et al. 2022). The odorant receptor (OR) chemoreceptor family, typically tuned to short-chain alcohols, ketones, and aromatic compounds, likely contributes toward anthropophilic host preference in A. aegypti (DeGennaro et al. 2013; McBride et al. 2014). In a complementary fashion, chemoreceptors from the ionotropic receptor (IR) family that are responsive to carboxylic acids and amines (Raji et al. 2019) and certain gustatory receptor (GR) family members that detect the volatile gas carbon dioxide (CO₂) (McMeniman et al. 2014) synergistically drive behavioral taxis of female mosquitoes toward human scent.

T2A-QF2 in-frame fusions have been applied to map expression patterns of target chemosensory genes from the OR, IR, and GR families in the olfactory appendages of Drosophila (Task et al. 2022), Anopheles coluzzii and Anopheles gambiae (Raji et al. 2019; Ye et al. 2022; Giraldo et al. 2024), and A. aegypti (Herre et al. 2022; Zhao et al. 2022). These neurogenetic tools leveraging QF2/QUAS-mediated binary expression (Riabinina et al. 2015) have also been used to express genetically encoded calcium indicators (GECIs) to study the ligand tuning dynamics of different sets of sensory neurons in the peripheral sensory organs of A. coluzzii and A. gambiae (Laursen et al. 2023; Raji et al. 2023; Giraldo et al. 2024) and taste neurons in the A. aegypti stylet (Jové et al. 2020). Application of T2A-QF2/QUAS-mediated cell-type-specific expression has also permitted optogenetic activation of CO₂-sensitive neurons in the maxillary palps of A. aegypti to help expand our understanding of CO₂-activated behavioral state changes in this mosquito (Sorrells et al. 2022). While optogenetics appears to have some success, there are limited reports of calcium imaging in the peripheral sensory organs in A. aegypti, likely because its olfactory appendages are enveloped by dark, highly melanized cuticle which dampens recording of stimulus-evoked fluorescent calcium signals from GECIs.

Moving centrally, the axonal processes of OSNs project to the primary olfactory processing center of the A. aegypti brain known as the antennal lobe (Ignell et al. 2005; Shankar and McMeniman 2020). In related insects such as Drosophila, olfactory information is locally processed and encoded in the antennal lobe via the action of excitatory and inhibitory local neurons (Chou et al. 2010; Yaksi and Wilson 2010; Hong and Wilson 2015), before being sent by projection neurons to higher-order brain centers involved in orchestrating innate and learned olfactory behaviors (Caron et al. 2013; Grabe et al. 2016; Dolan et al. 2019). Neurogenetic tools have allowed the recent application of GCaMP calcium indicators to study olfactory processing in the A. aegypti antennal lobe and other brain regions (Bui et al. 2019; Vinauger et al. 2019; Lahondère et al. 2020; Melo et al. 2020; Zhao et al. 2021, 2022; Wolff et al. 2023). These studies have facilitated identification of regions in the AL where human odor is putatively encoded by neurons expressing the OR co-receptor Orco (Zhao et al. 2022), as well as glomeruli that encode for flower odors (Lahondère et al. 2020) and oviposition cues (Melo et al. 2020). Since the antennal lobe is the first relay station for processing olfactory information, high-resolution mapping of the neuroanatomy of this brain center is crucial to discern the neural basis of mosquito olfactory coding.

Here, we describe a series of optimized genetic tools for neuroanatomical and functional mapping of the A. aegypti olfactory system. Using CRISPR–Cas9 homology-directed repair (HDR) donors that report the nature of transgene integration with a dual 3xP3-based marker system, we generate precise T2A-QF2 in-frame fusions in 3 target A. aegypti chemoreceptor genes, namely Ir8a, orco, and Gr1, for application in binary combination with various QUAS responders. Using Gr1+ OSNs as an example, we demonstrate that introgression of Gr1 T2A-QF2 driver and QUAS-GCaMP6s responder transgenes into a mutant yellow background that lacks cuticular melanization (Li et al. 2017) increases the signal obtained from peripheral sensory neurons during calcium imaging by 2–4-fold. We further show that Cre-loxP-mediated excision of commonly used 3xP3 markers (Berghammer et al. 1999) facilitates clear mapping of axonal projection patterns of these different subsets of OSNs into the antennal lobe, devoid of background marker interference. We provide an updated map of the A. aegypti antennal lobe detailing the projection patterns of OSNs expressing these chemoreceptors to specific glomeruli in this brain region. Finally, we demonstrate that neurons expressing Gr1 project to the largest glomerulus (MD1) in the A. aegypti antennal lobe and determine using calcium imaging with GCaMP6s (Chen et al. 2013) and the photoactivatable genetically encoded calcium indicator CaMPARI2 (Moeyaert et al. 2018) that this glomerulus mediates CO₂ detection. These optimized genetic tools will facilitate systematic studies that aim to decode the ligand tuning dynamics and principles of A. aegypti olfactory processing at both peripheral and central levels to improve our fundamental understanding of the chemosensory biology of this prolific disease vector.

Materials and methods

Mosquito strains and maintenance

The A. aegypti LVPib12 strain (Nene et al. 2007) was used as the genetic background for generation of all transgenic lines and subsequent assays. exu-Cas9 and yellow mutant strains were generated and provided by Li et al. (2017). Mosquitoes were maintained with a 12-h light:dark photoperiod at 27°C and 80% relative humidity using a standardized rearing protocol (Wohl and McMeniman 2023). All experiments were conducted with mated, nonblood fed A. aegypti females that were 5–10 days old. Adult mosquitoes were provided constant access to a 10% w/v sucrose solution. Stock and composite genotypes used in each figure panel are detailed in Supplementary Table 1 in Supplementary File 1.

Selection and in vitro transcription of single-guide RNAs

Single-guide RNA (sgRNA) target sites were identified in the coding sequences of orco (AAEL005776), Ir8a (AAEL002922), and Gr1 (AAEL002380) (Supplementary Table 2 in Supplementary File 1). Candidate sgRNAs at each locus were prioritized for downstream use based on their putative lack of off-target activity in the A. aegypti genome. sgRNAs were transcribed and purified according to the method of Kistler et al. (2015). Briefly, DNA templates for sgRNA synthesis were generated by PCR with 2 partially overlapping PAGE-purified oligos (IDT) for each target. sgRNA was subsequently produced using the MegaScript T7 in vitro transcription kit (Ambion) and purified using the MEGAclear transcription clean-up kit (Invitrogen). Prior to microinjection, sgRNA activity was confirmed by in vitro cleavage assays with purified recombinant Cas9 protein (PNA Bio, Inc., CP01-200) following the manufacturer's instructions.

T2A-QF2 donor constructs

A base T2A-QF2 donor construct (pBB) for CRISPR–Cas9-mediated homologous recombination into target chemoreceptor loci in A. aegypti was generated by sequential rounds of In-Fusion cloning (TakaraBio, 639650). This construct that was generated in a pSL1180 backbone has an EcoNI site for in-frame fusion of a 5′ homology arm from a target gene with T2A-QF2, and additional restriction sites including BssHII site for insertion of the 3′ homology arm. To survey for homology arms, genomic DNA regions spanning each target site were first PCR-amplified with CloneAmp (TakaraBio, 639298) using the following primers for orco (5′-TGCAAGTGGATCATTTGTCG-3′ and 5′-GTGCAATTGTGCCATTTTGA-3′), Ir8a (5′-CAAAGTATAATTTCGCCCCCTCC-3′ and 5′-CTCTATGGCAGCCAAGATATTGG-3′), and Gr1 (5′-AAGCCAGCTGGAAGGACATA-3′ and 5′-ACCGTTTGGAGGTTGAATTG-3′). PCR products were cloned into pCR2.1-TOPO (Invitrogen) for subsequent sequence verification. After determining the most common sequence clone for each region, homology arms flanking the CRISPR–Cas9 cut site were PCR-amplified and inserted into the pBB donor at the EcoNI site (5′ arm) and BssHII site (3′ arm) using the In-Fusion primers detailed in Supplementary Table 3 in Supplementary File 1, to generate a T2A in-frame fusion into the coding exon of interest. Three donor constructs that yielded successful integrations at these target loci included pBB-AaOrco, pBB-AaIR8a, and pBB-AaGr1. Each T2A-QF2 donor construct included a floxed 3xP3-DsRed2 transformation marker, as well as an unfloxed 3xP3-ECFP marker in the vector backbone outside the HDR cassette. This latter 3xP3-ECFP marker was used to assess putative vector backbone integration events at the target locus or alternate off-target integrations elsewhere in the genome. orco^QF2–3xP3 and Ir8a^QF2–3xP3 cassettes inserted in-frame as expected via canonical HDR. The Gr1^QF2–3xP3 cassette inserted in-frame but incorporated a duplicated copy of the donor cassette along with intervening plasmid backbone sequence downstream of the T2A-QF2 in-frame fusion via a noncanonical HDR repair event.

Mos1 mariner QUAS responder and germline Cre constructs

QUAS responder and germline Cre cassettes were generated by sequential rounds of In-Fusion cloning (TakaraBio) into template plasmid backbones for Mos1 mariner transposition (Coates et al. 1998). All QUAS reporter constructs included a 3xP3-ECFP transformation marker. The pMOS cassette for 15xQUAS-CaMPARI2 was modified to include a floxed 3xP3-ECFP marker, and the pMOS backbone for exu-Cre was modified to have a Polyubiquitin-EYFP marker using standard cloning methods. Final plasmids that yielded transformants included: pMosECFP-15xQUAS-mCD8GFP, pMosECFP-15xQUAS-GCaMP6s, pMos-loxP-ECFP-loxP-15xQUAS-CaMPARI2, and pMosEYFP-exu-Cre.

All template materials used to generate the constructs in this study are detailed in Supplementary Table 4 in Supplementary File 1. Stellar Competent Escherichia coli cells (Takara, 636763) were used for all cloning and plasmid preparations.

Generation of transgenic lines

T2A-QF2 knock-in lines were generated via CRISPR–Cas9-mediated HDR using embryonic microinjection. To generate the Gr1^QF2–3xP3 insertion, an injection mixture consisting of sgRNA (40 ng/µL), purified recombinant Cas9 protein (PNA Bio, 300 ng/µL), and donor plasmid (500 ng/µL) was prepared in microinjection buffer (5 mM KCl and 0.1 mM NaH₂PO₄, pH 7.2) and microinjected into the posterior pole of preblastoderm stage LVPib12 embryos at the Insect Transformation Facility at University of Maryland (UM-ITF) using standard methods. To generate the orco^QF2–3xP3 and Ir8a^QF2–3xP3 insertions, sgRNA (100 ng/µL) was mixed with the T2A-QF2 donor construct (100 ng/µL) for each target and microinjected into the posterior pole of transgenic A. aegypti preblastoderm stage embryos expressing Cas9 under the maternal germline promoter exuperantia (Li et al. 2017) at Johns Hopkins.

Transformed G₁ larvae from all knock-in lines were initially isolated via the visible expression of 3xP3-DsRed2 marker in eye tissue. Putative noncanonical HDR events were screened for by examining these G₁ larvae for co-expression of 3xP3-DsRed2 and 3xP3-ECFP in eye tissue. Transgenics were outcrossed to the LVPib12 wild-type (WT) line for at least 5 generations prior to attempting to generate homozygous strains. A single representative T2A-QF2 in-frame fusion event for each gene target was chosen for downstream characterization, prioritizing canonical HDR insertions for each locus if they were available. Precise insertion of each donor construct in these mosquito lines was confirmed by PCR amplification and subsequent Sanger sequencing of regions covering the homology arms and flanking sequences on either side of the insertion. For logistical feasibility of stock maintenance, redundant mosquito lines derived from noncanonical HDR events or canonical HDR events were discarded after successful establishment of these sequence-validated driver lines.

QUAS responder and exu-Cre strains were generated by co-injecting each pMOS donor construct (500 ng/µL) with a pKhsp82 helper plasmid (300 ng/µL) expressing the Mos1 transposase (Coates et al. 1998) for quasi-random integration into the genome. Embryo microinjections to generate these strains were carried out by UM-ITF using standard techniques. For QUAS responders, the G₁ offspring selected for line establishment were those that had the strongest 3xP3-ECFP marker expression levels in the eyes and ventral nerve cord, indicative of responder loci accessible for neuronal expression.

Cre-loxP-mediated excision of 3xP3 fluorescent markers

To remove floxed 3xP3 marker cassettes, we crossed males of each QF2 driver line (Ir8a^QF2–3xP3, orco^QF2–3xP3, Gr1^QF2–3xP3) to females of the exu-Cre line we generated. We then screened F₁ progeny for loss of the 3xP3 fluorescent markers. Precise excision was confirmed for all 3 driver lines by PCR and Sanger sequencing.

Mos1 mariner Splinkerette PCR

QUAS and exu-Cre transgenes inserted via Mos1 mariner transposition were mapped to chromosomal locations (AaegL5.0 genome assembly) using a modified Splinkerette PCR (Potter and Luo 2010). Genomic DNA from single transgenic individuals was digested using the restriction enzymes BamHI-HF, BglII, and BstYI (New England BioLabs) in separate reactions. Digests were left overnight (∼16 h). BstYI reactions were subsequently heat-inactivated at 80°C for 20 min according to the recommended protocol. BamHI reactions were purified using the QIAquick PCR Purification Kit (QIAgen) according to manufacturer instructions and eluted in 50 μl H₂O after 4 min of incubation at 50°C.

Digests of genomic DNA were ligated to annealed Splinkerette (SPLNK) oligos as described (Potter and Luo 2010). SPLNK oligonucleotides 5′-GATCCCACTAGTGTCGACACCAGTCTCTAATTTTTTTTTTCAAAAAAA-3′ and 5′-CGAAGAGTAACCGTTGCTAGGAGAGACCGTGGCTGAATGAGACTGGTGTCGACACTAGTGG-3′ were first annealed and ligated to digested genomic DNA. The first- and second-round PCR amplification steps were modified using the standard SPLNK oligos and new primers designed for the inverted repeat regions of the Mos1 mariner transposon. PCR products were amplified using Phusion High-Fidelity DNA Polymerase (NEB).

The first-round splinkerette PCR was carried out using the primers 5′-CGAAGAGTAACCGTTGCTAGGAGAGACC-3′ and 5′-TCAGAGAAAACGACCGGAAT-3′ for the right inverted repeat and 5′-CGAAGAGTAACCGTTGCTAGGAGAGACC-3′ and 5′-CACCACTTTTGAAGCGTTGA-3′ for the left inverted repeat. The second-round splinkerette PCR was carried out using the primers 5′-GTGGCTGAATGAGACTGGTGTCGAC-3′ and 5′-TCCGATTACCACCTATTCGC-3′ for the right inverted repeat and 5′-GTGGCTGAATGAGACTGGTGTCGAC-3′ and 5′-ATACTGTCCGCGTTTGCTCT-3′ for the left inverted repeat. For QUAS-CaMPARI2, the extension time of the second-round PCR was lengthened to 4 min to amplify longer segments of flanking DNA. PCR products were gel purified and Sanger sequenced with additional sequencing primers for the right (5′-AAAAATGGCTCGATGAATGG-3′) and left (5′-GGTGGTTCGACAGTCAAGGT-3′) inverted repeats. BLAST searches were used to map splinkerette fragments derived from each Mos1 mariner cassette to coordinate locations in the genome at canonical TA dinucleotides (Richardson et al. 2009), and insertion sites (Supplementary Table 5 in Supplementary File 1) were subsequently confirmed by PCR.

Genotyping Gr1^QF2–3xP3 and Gr1^QF2-MF

Gr1^QF2–3xP3 and Gr1^QF2-MF knock-ins were genotyped using a multiprimer PCR assay with the forward primer: 5′-CATGTACATCCGCAAGTTGG-3′ and 2 standard reverse primers: 5′-TGTTAGTGAGATCAGCGAACCT-3′ and 5′-GATCAACCCACAGATGACGA-3′. Fragments for size-based genotyping were amplified via DreamTaq (Thermo Scientific) and analyzed by conventional agarose gel electrophoresis. Each of the reverse primers was used at half the normal concentration. This resulted in a single 689-bp amplicon in homozygous mosquitoes; a single 884-bp amplicon in WT mosquitoes; and 2 amplicons, 1 at 689 bp and 1 at 884 bp, in heterozygous mosquitoes.

To characterize the nature of the HDR insertion in Gr1^QF2–3xP3 which contained both 3xP3-DsRed2 and 3xP3-ECFP markers indicative of both donor cassette and plasmid backbone integration, we used PCR to amplify 2 overlapping fragments. The first amplicon (6,770 bp) had primers anchored in the genomic region outside the left homology arm (5′-TCGCTGAGTGATGAGGGTTT-3′) and within ECFP (5′-CTTCTCGTTGGGGTCTTTGC-3′). The second amplicon (8,432 bp) had primers anchored within ECFP (5′-GAGGAGCTGTTCACCGGG-3′) and in the genomic region outside the right homology arm (5′-CATGAATGCCCAAGACCATCT-3′). PCR fragments were gel purified (Wizard SV Gel and PCR Cleanup System, Promega) and Nanopore sequenced (Plasmidsaurus) for assembly.

Genotyping Ir8a^QF2–3xP3 and Ir8a^QF2-MF

Ir8a^QF2–3xP3 and Ir8a^QF2-MF knock-ins were genotyped using a multiprimer PCR assay with the forward primer: 5′-AGGAGATTGCGCTTGTCCTA-3′ and 2 reverse primers: 5′-CCCCGACATAGTTGAGCATT-3′ and 5′-TGTTAGTGAGATCAGCGAACCT-3′. Each of the reverse primers was used at half the normal concentration. This resulted in a single 560-bp amplicon in homozygous mosquitoes; a single 501-bp amplicon in WT mosquitoes; and 2 amplicons, 1 at 560 bp and 1 at 501 bp in heterozygous mosquitoes.

Genotyping orco^QF2–3xP3 and orco^QF2-MF

orco^QF2–3xP3 and orco^QF2-MF knock-ins were genotyped using conventional PCR. The PCR used the forward primer: 5′-GCGATAGCGTCAAAAACGTA-3′ and reverse primer: 5′-ATTCCTTGAAGGTCCATTGCAG-3′. This resulted in a 1,842-bp amplicon corresponding to the orco^QF2-MF allele, a 3,129-bp amplicon corresponding to the orco^QF2–3xP3 allele, and a 367-bp amplicon corresponding to the WT allele. Heterozygotes had both WT and transgenic PCR bands.

Genotyping 15xQUAS-mCD8::GFP

15xQUAS-mCD8::GFP was genotyped using conventional PCR. The PCR used the forward primer: 5′-TCCAGCCGATAGGAACAATC-3′ and reverse primer: 5′-CAAATCCGAATTTCCCGTAA-3′. This resulted in a single 5,797-bp amplicon for homozygotes and a 444-bp amplicon for the WT allele. Heterozygotes typically only had the WT PCR band given the size differential between these 2 amplicons.

Genotyping 15xQUAS-GCaMP6s

15xQUAS-GCaMP6s was genotyped using a multiprimer PCR with the forward primer: 5′-CCAATCCCTCCAAAACAAGA-3′; and 2 reverse primers: 5′-ACGCTTTCGACAGATTCGTT-3′ and 5′-CACCACTTTTGAAGCGTTGA-3′. Each of the reverse primers was used at half the normal concentration. This resulted in a single 529-bp amplicon in homozygous mosquitoes; a single 379-bp amplicon in WT mosquitoes; and 2 amplicons, 1 at 529 bp and 1 at 379 bp in heterozygous mosquitoes.

Genotyping 15xQUAS-CaMPARI2

15xQUAS-CaMPARI2 was genotyped using a multiprimer PCR assay with the forward primer: 5′-GTTTGACCAAATGCCGTTTC-3′ and 2 standard reverse primers: 5′-GTCGATAGGCGCGTAGTGTA-3′ and 5′-CACCACTTTTGAAGCGTTGA-3′. Each of the reverse primers was used at half the normal concentration. This resulted in a single 645-bp amplicon in homozygous mosquitoes; a single 874-bp amplicon in WT mosquitoes; and 2 amplicons, 1 at 645 bp and 1 at 874 bp in heterozygous mosquitoes.

Transgenic stock maintenance and composite genotypes

Gr1^QF2–3xP3, Gr1^QF2-MF, Ir8a^QF2–3xP3, and Ir8a^QF2-MF driver lines were maintained as homozygous stocks. orco^QF2–3xP3 was maintained as a heterozygous stock by outcrossing to LVPib12 each generation. orco^QF2-MF was maintained as a heterozygous stock by outcrossing to either LVPib12 or QUAS-mCD8::GFP each generation and screening for GFP fluorescence in olfactory tissues of the progeny. 15xQUAS-mCD8::GFP and 15xQUAS-CaMPARI2 responder lines were maintained as homozygous stocks. The exu-Cre line was maintained as a heterozygous stock by outcrossing to LVPib12 each generation. For long-term maintenance, we have determined that all driver and responder stocks except orco^QF2–3xP3 and exu-Cre are capable of being maintained as homozygous stocks. We do not observe any major fitness effects that preclude laboratory rearing of these stocks when maintained as indicated.

Immunohistochemistry

Immunostaining of female A. aegypti brains was performed as previously described (Shankar and McMeniman 2020), with minor modifications. Briefly, severed mosquito heads were fixed in 4% paraformaldehyde (Milonig's buffer, pH 7.2) for 3 h after which brains were carefully dissociated from the head capsule, pigmented ommatidia, and air sacs. Dissected brains were then subjected to three 20-min washes at room temperature in PBST (0.1 M PBS with 0.25% Triton-X 100) and allowed to incubate overnight in a blocking solution consisting of 2% normal goat serum (NGS) and 4% Triton-X 100 in 0.1 M PBS at 4°C. Brains were then washed 3 times for 20 min each in PBST and incubated for 3 days at 4°C in a primary antibody solution containing mouse anti-BRP (DSHB, nc82-s, AB_2314866, 1:50 v/v) targeting the presynaptic active zone protein Bruchpilot (Hofbauer et al. 2009) and rabbit anti-GFP (Invitrogen, A-6455, 1:100 v/v) targeting mCD8::GFP. Brains were then washed 3 times for 20 min each in PBST and incubated for 3 days at 4°C in a secondary antibody solution consisting of goat antimouse Cy3 (Jackson ImmunoResearch, AB_2338680, 1:200 v/v) and goat antirabbit Alexa Fluor 488 (Invitrogen, A-11008, 1:200 v/v). All primary and secondary antibody dilutions were prepared in PBST with 2% v/v NGS. Brains were finally washed 3 times for 20 min each in PBST at room temperature and mounted in 20 µL of SlowFade Gold antifade mountant (Invitrogen, S36936) on glass slides with coverslip bridges (number 2–170 μm).

Immunohistochemistry image acquisition settings

Brain immunostaining images were acquired on a single-point laser scanning Carl-Zeiss LSM 780 confocal microscope. To capture images of the entire adult brain, a 10× objective lens (0.3 NA, Plan-Apochromat) was used. Excitation of Cy3 signal was achieved with a 561-nm solid-state laser line at 0.05% laser power and GaAsP detector gain set to 825. A 488-nm laser line was used to excite Alexa Fluor 488 (20% laser power, detector gain at 825). We additionally acquired images with a 20× objective lens (0.8 NA, Plan-Apochromat) to perform 3D reconstructions of the antennal lobes. For these, the power of the 488-nm laser line was adjusted to 5%. For each antennal lobe, 60 z-slices with a z-step size of 1 μm and a 1,024 × 1,024-pixel resolution were acquired.

Antennal lobe reconstructions

Three-dimensional morphological reconstructions of left antennal lobes were performed as previously described (Shankar and McMeniman 2020). Briefly, confocal images were imported into Amira (FEI Houston Inc.) and then segmented by highlighting all pixels across a z-stack occupied by individual glomeruli. The nc82 (Bruchpilot) channel was used for manual segmentation of individual glomeruli. The GFP channel was then used to identify orco+, Ir8a+, and Gr1+ glomeruli. Cross-referencing signals obtained from nc82 and GFP channels within and between samples in this dataset helped to clearly delineate glomerular boundaries. 3D and 2D antennal lobe models were generated by surface rendering. The number of GFP-labeled glomeruli in 3 replicate left antennal lobe reconstructions per genotype from orco^QF2-MF > 30xQUAS-mCD8::GFP, Ir8a^QF2-MF > 30xQUAS-mCD8::GFP, and Gr1^QF2-MF > 30xQUAS-mCD8::GFP females was counted. The total number of glomeruli per lobe was counted in 7 of these samples: orco^QF2-MF > 30xQUAS-mCD8::GFP (n = 3), Ir8a^QF2-MF > 30xQUAS-mCD8::GFP (n = 3), and Gr1^QF2-MF > 30xQUAS-mCD8::GFP (n = 1).

Glomerular volume and frequency

Glomerular volumes were obtained from the left antennal lobe using the nc82 channel. To name glomeruli, we first identified landmark glomeruli in each antennal lobe sample using a systematic A. aegypti antennal lobe reference key (Shankar and McMeniman 2020), as this updated map better reflected glomerular organization in our dataset relative to a previous map (Ignell et al. 2005). Each antennal lobe glomerulus labeled with GFP was named based on its spatial position relative to these landmarks and flanking glomeruli. We classified glomeruli as spatially “invariant” or “variant” based on their frequency of identification. A threshold frequency of 80% or more across reconstructions was designated for the classification of spatially invariant glomeruli. Glomerular volume and frequency values were calculated from a pooled dataset consisting of the 7 reconstructions where we named all constitutive glomeruli in the same left antennal lobe samples used for glomerular counts.

Confocal imaging of peripheral olfactory appendages

Live antenna, palp, and proboscis tissue were dissected in 0.1 M PBS and immediately mounted in SlowFade Gold antifade mountant (Invitrogen, S36936). Images were acquired on a Carl-Zeiss LSM 780 confocal microscope within 1 h of dissection. To excite the GFP signal, the 488-nm laser line was used at 5% laser power. An additional DIC channel was used to visualize gross morphology of the peripheral tissue. Images of the antennae were acquired with a 20× objective lens (0.8 NA, Plan-Apochromat), while images of the palp and labella of the proboscis were taken with a 40× (1.3 NA, Plan-Apochromat) oil immersion objective.

Mosquito preparation for peripheral calcium imaging

Five- to 10-day-old female mosquitoes were transferred to plastic fly vials (Flystuff, 32–110) containing a cotton ball with distilled water for fasting (1 mosquito per vial). The vials were placed in a climate-controlled incubator (27°C, 80% relative humidity) for 20–24 h before testing. Mosquitoes were cold anesthetized for 3 min, and the wings and legs were removed. They were then placed on a 3D-printed holder that allowed for the mosquito to rest upside down, exposing the ventral side of the palps where the CO₂ sensing neurons are found. The dorsal side of the palps was placed on double-sided tape (Scotch, 137DM-2) to fix them in position. The proboscis was taped down to reduce movement of the preparation.

Mosquito preparation for antennal lobe recordings

Five- to 10-day-old female mosquitoes were fasted as described above. Females were briefly knocked out on ice, and legs and wings were removed. They were then situated within a small oval cut into tin foil that was just big enough so that the top of the head and thorax rested above the tin foil and the olfactory organs and rest of the body were below and free to move. UV glue (BONDIC CECOMINOD032561) was applied with a thin tungsten wire to secure the upper thorax and head to the tin foil. Next, calcium-free saline (150 mM NaCl, 3.4 mM KCl, 5 mM glucose, 1.8 mM NaHCO₃, 1 mM MgCl₂, 25 mM HEPES; pH 7.1; 0.22 µm filtered sterilized) was added to cover the top of the head and a 31-gauge needle (BD 328289) was used to cut a square through the cuticle from the base of the pedicels to the vertex. The cuticle was gently removed with sharp forceps, and calcium-free saline was then replaced with saline with calcium (same saline as above with 1.7 mM CaCl₂) by removal with a Kimwipe and simultaneous addition of new saline with a pipette at which point imaging commenced.

CO₂ delivery for calcium imaging

5% CO₂ (X02AI95C2000117, Airgas) was mixed with clean air (UZ300, Airgas) using mass flow controllers (MC-series 100, MC-series 500, Alicat Scientific) to obtain the desired concentration at a flow of 2 mL/s. The stimulus was controlled by a solenoid valve (ETO-3-12, Clippard) and valve driver (Automate Scientific, ValveLink 8.2) and brought into a carrier flow of humidified air flowing at 6 mL/s for a total flow of 8 mL/s. A compensatory clean air flow was on at 2 mL/s whenever the stimulus was off to ensure a constant flow of 8 mL/s.

Calcium imaging system and analysis

Calcium signals were measured using an epifluorescence microscope (BX51W1, Olympus) and an LED light source (Lambda HPX-L5, Sutter Instrument) at 50× magnification for peripheral recordings and 20× for AL recordings. Recordings were carried out at 30 fps using a CMOS camera (Orca-Fusion C14440, Hamamatsu Photonics K.K.) controlled by micro-manager. The videos were analyzed using FIJI, and the data were analyzed using MATLAB R2018b (The MathWorks, Inc.).

Live mosquito preparation for CaMPARI2 photoconversion

To prepare mosquitoes for CaMPARI2 photoconversion (Moeyaert et al. 2018), mosquitoes were cold anesthetized and tethered to an imaging chamber. To do this, the thorax of a female mosquito was first affixed to the ventral surface of a 35-mm petri dish lid (Eppendorf, 0030700112) using UV-curing adhesive (Bondic) immediately next to a 15-mm-diameter circular hole made in the lid center. Two additional drops of adhesive were applied to the ommatidia on the extremities of the mosquito head to prevent head movement. A small piece of clear tape (Duck EZ Start, Heavy Duty Packaging Tape) was then adhered over the center hole. The dorsal surface of the mosquito head was then gently affixed to the ventral adhesive tape surface covering the hole. An excised section of plastic coverslip (5 mm × 3 mm) was then affixed to the tape and used to shield the antennae from the adhesive tape surface and suspend these sensory appendages in the air.

The imaging chamber with head-fixed mosquito was then inverted, and a rectangular incision ∼400 µm × 200 µm was cut through the tape window where the dorsal head cuticle and ommatidia were affixed. The wide boundary of the incision was typically made immediately adjacent to the first antennal subsegment along the lateral–medial brain axis, while the short boundary of the incision extended along the dorsal–ventral brain axis. To create this window, segments of ommatidia and bridge cuticle between the left and right eyes were gently cut and excised using a surgical stab knife (Surgical Specialties Corporation, Sharpoint, Part # 1038016) to reveal the underlying antennal lobes. The exposed antennal lobes were then immediately immersed in an A. aegypti Ringer's solution (Beyenbach and Masia 2002) composed of 150 mM NaCl, 3.4 mM KCl, 5 mM glucose, 1.8 mM NaHCO₃, 1 mM MgCl₂, 25 mM HEPES, and 1.7 mM CaCl₂; pH 7.1. Mosquitoes were allowed to recover for a period of 15 min from cold anesthesia and surgery in a humidified chamber at room temperature prior to imaging.

CaMPARI2 photoconversion

For CaMPARI2 photoconversion, the tethered preparation was placed under a 20× water dipping objective (Olympus XLUMPLFLN20XW, 1.0 NA), ensuring that the antennal lobes expressing basal green CaMPARI2 signal were in focus. Each preparation was exposed to a combined photoconversion-odor stimulation regime consisting of repetitive duty cycles of four 500-ms pulses of 405-nm light from an LED driver (Thorlabs, DC4104, 1,000 mA current setting) synchronized with a 1-s odorant pulse as previously outlined (Fosque et al. 2015), for 75 cycles with a total protocol duration of ∼41 min.

Odorant delivery for CaMPARI2 photoconversion

A 1-mL/s stream of 5% CO₂ was diluted 1:5 into the carrier airstream for a final concentration at the specimen of 1% CO₂ and was then piped via Teflon tubing into a carrier airstream of humidified synthetic air that was directed at the olfactory appendages of the mosquito using a plastic pipette. During CaMPARI2 photoconversion assays, the tethered mosquito preparation always received a constant amount of airflow (5 mL/s) during odor onset/offset from the stimulus pipette. This was achieved via solenoid valves simultaneously switching or combining humidified synthetic air, 5% CO₂ (Airgas).

CaMPARI2 sample processing

Following photoconversion, the mosquito was gently untethered from the imaging chamber and the head severed and fixed in Milonig's buffer for 20 min. The brain was then dissected out in calcium-free Ringer's solution composed of 150 mM NaCl, 3.4 mM KCl, 5 mM glucose, 1.8 mM NaHCO₃, 1 mM MgCl₂, 25 mM HEPES, and 10 mM EGTA. To stain glomerular boundaries, we incubated each brain in Alexa Fluor 647 Phalloidin (Invitrogen, A22287) prepared in calcium-free Ringer's solution (1:40 v/v dilution) for 30 min. To prepare Alexa Fluor 647 phalloidin for use in imaging, first, a 400× DMSO stock solution was prepared according to the manufacturer's instructions by dissolving the fluorophore in 150 µL of DMSO. One microliter of this DMSO stock was diluted in 399 µL calcium-free Ringer's solution to yield a 1× stock. This 1× stock was then further diluted 1:40 in calcium-free Ringer's solution for staining. Brains were transferred directly from this solution into 20 µL of SlowFade Gold antifade mountant (Invitrogen, S36936) on glass slides with coverslip bridges (number 2–170 μm) for CaMPARI2 and phalloidin imaging.

CaMPARI2 image acquisition settings

Antennal lobes from CaMPARI2 photoconversion assays were imaged with a 63× (1.4 NA) oil immersion objective on a Zeiss 880, Airyscan FAST super-resolution single-point scanning microscope. Excitation of red CaMPARI2 signal was achieved with a 561-nm solid-state laser line at 14% laser power. Green CaMPARI2 was excited with a 488-nm argon laser line at 10% laser power. To visualize glomerular boundaries, a 633-nm diode laser was used to excite the Alexa-647 phalloidin fluorophore at 40% laser power. Master detector gain was set to a value of 800. We captured 0.987 μm z-slices of 1,572×1,572-pixel resolution in the FAST mode. Raw images were further processed by applying the Airyscan method with “auto” processing strength.

CaMPARI2 image analysis

Image analysis was carried out in Fiji (http://imagej.net/Fiji). We first applied a median filter (radius = 2 pixels) to remove noise and then a rolling ball subtraction (rolling ball radius = 80 pixels) to correct for nonuniformity of background intensities. ROIs were defined by manually segmenting the MD1 glomerulus using the freehand selection tool. The integrated density (mean gray value × area) for all z-slices of the ROI, which included all representative slices of a target glomerulus, was calculated in the green (488 nm) and red (560 nm) imaging channels. The final measure of photoconversion, the red-to-green ratio photoconversion (R/G), was calculated as:

$$\begin{array}{rl}R/G& =\mathrm{Average}\mathrm{integrated}\mathrm{density}\mathrm{of}\mathrm{RO}{\mathrm{I}}^{(\mathrm{RED})}\\ & /\mathrm{Average}\mathrm{integrated}\mathrm{density}\mathrm{of}\mathrm{RO}{\mathrm{I}}^{(\mathrm{GREEN})}\end{array}$$

Statistical analysis

We tested normality of raw data with a Kolmogorov–Smirnov test. To test for the statistical significance of differences observed in different experiments, dependent on context we used Wilcoxon rank-sum, Mann–Whitney U and paired t-tests, and ANOVA with post hoc comparisons. Statistical analyses were performed in MATLAB R2018b (The MathWorks, Inc.) and GraphPad Prism Software version 8.4.0. Details of statistical methods are reported in the figure legends.

Results

Targeted genetic access to A. aegypti OSN populations using CRISPR–Cas9 HDR donors that report the nature of transgene integration

CRISPR–Cas9-mediated T2A in-frame fusions (Diao and White 2012) are a broadly used genetic strategy to capture endogenous expression patterns of target genes of interest in varied organisms, including A. aegypti, where this technology is routinely applied to map the expression patterns of hygrosensors, thermoreceptors, chemoreceptors, and other neural genes (Matthews et al. 2019; Jové et al. 2020; Zhao et al. 2021, 2022; Herre et al. 2022; Laursen et al. 2023). Of note, all existing T2A-QF2 integrations reported to date in A. aegypti have employed 3xP3 fluorescent transgenesis markers (Berghammer et al. 1999) to identify transformants. With current designs employed in A. aegypti, these 3xP3 transgenesis markers cannot be easily removed post-integration due to a lack of specific sequence motifs flanking these cassettes to facilitate their excision. Furthermore, current donor construct designs used for CRISPR–Cas9-mediated HDR at target loci in A. aegypti do not report the nature of cassette integration.

Specifically, 2 common integration outcomes may occur in the presence of a double-stranded break and circular homologous donor plasmid during CRISPR–Cas9-mediated HDR across various species. These include canonical HDR repair events, as well as noncanonical insertions in which the intended insertion is duplicated at the target site, with the duplicate copies separated by intervening plasmid sequence. This latter phenomenon has been observed in different organisms including Drosophila (Bier et al. 2018; Zirin et al. 2022; Loehlin et al. 2023). Within the context of T2A in-frame fusions, canonical HDR repair events result in precise integration of a single copy of the T2A-QF2 donor cassette into the target site. On the other hand, noncanonical duplications, while also yielding a precise integration of T2A-QF2 into the target site as intended, may complicate genotyping of lines post-establishment as the plasmid backbone and a duplicate copy of the donor cassette are also integrated.

We therefore updated the design of our circular homologous recombination construct used for T2A-QF2 in-frame fusions in A. aegypti to include a T2A-QF2 cassette with a 3xP3-DsRed2 eye marker flanked by loxP sites (floxed), as well as a second nonfloxed 3xP3-ECFP eye marker in the vector backbone (Fig. 1a). This construct design facilitates the rapid identification of transgene integrations generated by canonical HDR (i.e. those with DsRed2 positive eyes only) and those that also possibly incorporate plasmid backbone indicative of a multiplexed integration event of the donor cassette (i.e. those with DsRed2- and ECFP-positive eyes). In this donor construct design, loxP sites flanking the 3xP3-DsRed2 eye marker also enable marker removal from canonical integration events using Cre-loxP excision, which is highly efficient in A. aegypti (Häcker et al. 2017). Similarly, duplication of the floxed 3xP3-DsRed2 donor cassette and insertion of the intervening plasmid backbone sequence, including the 3xP3-ECFP marker, from multiplexed noncanonical recombination events can be resolved to yield an integration reflective of canonical HDR using this excision method.

Fig. 1.

A dual transgenesis marker system reports canonical and noncanonical HDR repair events during CRISPR–Cas9-facilitated T2A-QF2 in-frame fusions in A. aegypti chemoreceptor genes. a) Schematic of circular donor plasmid used for CRISPR–Cas9-mediated HDR to generate T2A-QF2 in-frame fusion driver lines for the A. aegypti orco, Ir8a, and Gr1 genes. The targeting construct contains a 3xP3-DsRed2 transgene (transgenesis marker 1) flanked by loxP sites in the intended integration cassette and an additional 3xP3-ECFP transgene (transgenesis marker 2) positioned in the vector backbone. LHA, left homology arm; RHA, right homology arm; gRNA target site for each gene indicated by arrows. b) Marker expression in the head of fourth-instar larvae from orco^QF2–3xP3, Ir8a^QF2–3xP3, and Gr1^QF2–3xP3 driver lines. Heads outlined with dashed lines, arrows indicate marker expression in larval eyes c, d) Schematics of the canonical HDR integration events that generated the orco^QF2–3xP3 and Ir8a^QF2–3xP3 T2A-QF2 driver lines which are marked by only a single 3xP3-DsRed2 transgene flanked by loxP sites. e) Schematic of the noncanonical integration event that generated the Gr1^QF2–3xP3 driver line, marked by 2 3xP3-DsRed2 transgenes flanked by loxP sites, separated by intervening vector sequence with a 3xP3-ECFP transgene. The donor cassette is duplicated. Homology arms are shaded, and the positioning of loxP sites is indicated with arrows in a) and c—e).

To generate transgenic A. aegypti chemoreceptor-QF2 driver lines using this updated targeting strategy, we first designed gRNAs and circular HDR donor constructs to integrate T2A-QF2 in-frame into the coding exons of 2 olfactory co-receptor genes: orco and Ir8a which have been targeted previously using alternative construct designs (Herre et al. 2022; Zhao et al. 2022), as well as the CO₂ receptor complex gene Gr1 (McMeniman et al. 2014) which has not been targeted before (Fig. 1a). Using this strategy, QF2 was precisely integrated in-frame into Exon 3 of each target gene, placing the expression of this transcription factor under control of the endogenous regulatory elements for each locus. Using this approach, we successfully recovered precise T2A in-frame fusion integrations in orco and Ir8a via canonical HDR as indicated by the presence of a 3xP3-DsRed2 marker only in these lines (Fig. 1b–d). We also obtained a precise T2A in-frame fusion integration event in Gr1 that included 2 copies of the T2A-QF2 donor cassette and intervening plasmid backbone as indicated by the presence of both 3xP3-DsRed2 and 3xP3-ECFP markers in transformants (Fig. 1b and e). The orientation of this latter insertion in the A. aegypti genome was verified using long-read Nanopore sequencing, and we further confirmed that this sequence orientation was not found in the original donor plasmid using the same method and therefore was the result of noncanonical HDR (Fig. 1e).

Of note during isolation of the above T2A-QF2 lines, in the G₁ generation post-microinjection we observed only noncanonical HDR events for Gr1, a mixture of canonical and noncanonical HDR events for Ir8a, and a single canonical HDR event for orco (Supplementary Table 6 in Supplementary File 1). These differing HDR repair outcomes were also observed across different formats of Cas9 delivery and microinjection suppliers (Supplementary Table 6 in Supplementary File 1). We proceeded to characterize a single noncanonical HDR event carrying both 3xP3-DsRed2 and 3xP3-ECFP transgenesis markers for the Gr1 integration as that was the only option. Given their availability, single representative lines were selected for downstream characterization for Ir8a and orco integrations which each solely carried a 3xP3-DsRed2 transgenesis marker.

Each of these 3xP3-marked chemoreceptor-QF2 driver lines (orco^QF2–3xP3, Ir8a^QF2–3xP3, and Gr1^QF2–3xP3) was crossed with a 15xQUAS-mCD8::GFP responder strain that we generated via Mos1 mariner transposition (Coates et al. 1998; Supplementary Table 7 in Supplementary File 1). In contrast to the only other currently available QUAS-mCD8::GFP responder strain for this species, which includes a SV40 terminator for the mCD8::GFP gene (Matthews et al. 2019; Herre et al. 2022), we flanked this fluorescent reporter with both Syn21 and p10 3′UTR sequences, shown to act as translational enhancers in Drosophila (Pfeiffer et al. 2012) to generate a line that robustly labels neuronal membrane and processes in this mosquito. Olfactory tissues from the F₁ progeny of these crosses were then surveyed for the presence of membrane-tethered GFP in OSNs. Confocal analyses of adult peripheral sensory appendages revealed strong GFP labeling of orco + OSN dendrites and cell bodies on the antenna, maxillary palp and labella of the proboscis of orco^QF2–3xP3 > 15xQUAS-mCD8::GFP females (Fig. 2a). Strong labeling of Ir8a + OSNs within the antennal flagella (Fig. 2b) of Ir8a^QF2–3xP3 > 15xQUAS-mCD8::GFP and that of Gr1+ OSNs in maxillary palp tissue (Fig. 2c) of Gr1^QF2–3xP3 > 15xQUAS-mCD8::GFP females were also detected.

Fig. 2.

T2A-QF2 in-frame fusions facilitate visualization of orco+, Ir8a+ and Gr1+ OSNs on the olfactory appendages of A. aegypti. OSNs were labeled with membrane-tethered GFP (mCD8::GFP) in the: a) antennae (left), maxillary palps (middle), and labella of the proboscis (right) of orco^QF2–3xP3 > 15xQUAS-mCD8::GFP females. b) Antennae of Ir8a^QF2–3xP3 > 15xQUAS-mCD8::GFP females, and c) maxillary palps of Gr1^QF2–3xP3 > 15xQUAS-mCD8::GFP females. A schematic of the mosquito head with organs exhibiting observable GFP fluorescence for each line is shown. Scale bars: 50 µm.

Expression patterns from these T2A-QF2 knock-ins were consistent with a previous A. aegypti LVPib12 strain neurotranscriptome analysis using bulk RNAseq (Matthews et al. 2016) that revealed broad orco expression across adult olfactory tissues, along with strong, tissue-specific expression of Ir8a and Gr1 in antennal and maxillary palp tissue, respectively. The peripheral GFP labeling patterns that we observed for our orco^QF2–3xP3 and Ir8a^QF2–3xP3 lines developed here were also consistent with those observed for other T2A-QF2 in-frame fusions recently generated in the last coding exon of these genes (Herre et al. 2022). The pattern of GFP labeling that we observed for our Gr1^QF2–3xP3 driver line also mirrored that of another T2A in-frame fusion in the last coding exon of Gr3 (Herre et al. 2022), with both lines labeling OSNs localized to capitate peg sensilla in the A. aegypti maxillary palps.

The complementary transgenic lines developed here thus expand the toolkit to genetically access and label target A. aegypti OSN populations. Using these new T2A-QF2 driver lines and our 15xQUAS-mCD8::GFP responder line equipped with translational enhancers, we conclude that at the resolution of transgenic labeling and confocal imaging performed here, orco+ neurons clearly innervate all 3 olfactory sensory appendages in A. aegypti, whereas Ir8a+ and Gr1+ OSNs strongly innervate the antennae and maxillary palps, respectively.

A mutant A. aegypti strain with reduced cuticular pigmentation enables sensitive transcuticular imaging of OSN activity

To initially validate the applicability of these T2A-QF2 in-frame fusions for functional imaging of neural activity in A. aegypti, we first performed peripheral calcium imaging from CO₂ neurons on the ventral surface of the maxillary palps. The maxillary palps are the anatomical site of CO₂ detection in this species (Kellogg 1970; Grant et al. 1995), where the dendrites and underlying cell bodies of capitate peg A (cpA) neurons that co-express the CO₂ receptor complex subunits Gr1, Gr2, and Gr3 are localized (Jones et al. 2007; Lu et al. 2007).

To do this, we crossed our Gr1^QF2–3xP3 driver with a 15xQUAS-Syn21-GCaMP6s-p10 responder strain (15xQUAS-GCaMP6s) that we engineered via Mos1 mariner transposition (Supplementary Table 7 in Supplementary File 1), and imaged maxillary palp tissue in F₁ progeny carrying both transgenes. Although peripheral calcium imaging has been successfully used to study stimulus-evoked activity in A. coluzzii and A. gambiae antenna and maxillary palps (Afify et al. 2019; Laursen et al. 2023; Raji et al. 2023; Giraldo et al. 2024) and responses to tastants in the dissected stylet of A. aegypti (Jové et al. 2020), we observed that the darker pigmentation of the A. aegypti cuticle posed a major challenge toward measuring changes in GCaMP6s fluorescence in response to stimulation with CO₂. In particular, we hypothesized that this cuticular barrier optically occluded and dampened odor-evoked changes in fluorescence.

As cuticle covering all sensory appendages of WT A. aegypti including the maxillary palps is highly melanized, we sought to lighten cuticular pigmentation by introgressing both our Gr1^QF2–3xP3 and 15xQUAS-GCaMP6s transgenes into a yellow mutant background (Li et al. 2017). This was performed as the yellow gene is important for cuticular melanization in various insect species (Biessmann 1985; Wittkopp et al. 2002; Berni et al. 2022; Gong et al. 2024), and mutants of this gene in A. aegypti (Li et al. 2017) have lighter cuticle (Fig. 3a). We then compared odor-evoked changes in GCaMP6s fluorescence by stimulating Gr1^QF2–3xP3 > 15xQUAS-GCaMP6s female mosquitoes in both yellow mutant (y¹) and WT genetic backgrounds in response to stimulation with various concentrations of CO₂.

Fig. 3.

Aedes aegypti cuticular pigmentation mutant enables peripheral calcium imaging of CO₂-evoked activity from Gr1+ OSNs on the maxillary palps with enhanced sensitivity. a) Female A. aegypti of WT (left panel) and yellow mutant [y¹] (right panel) backgrounds. b) GCaMP6s fluorescence in maxillary palp neurons before (left panel) and during 1% CO₂ stimulation (right panel). c) GCaMP6s traces of CO₂-sensitive neurons in the maxillary palps after a 1% CO₂ pulse for 1 s (bar) from WT (Gr1^QF2–3xP3 > QUAS-GCaMP6s) and yellow mutant (Gr1^QF2–3xP3 [y¹] > QUAS-GCaMP6s [y¹]) females. The solid line represents the mean and the shaded area the standard error of the mean (n = 10 WT, n = 11 yellow). d) Maximum change in fluorescence to increasing concentrations of 1-s CO₂ pulses in WT and y¹ females. The dots represent the mean, and the error bars the standard error of the mean (n = 10 WT, n = 11 y¹). Wilcoxon rank-sum test, *P < 0.05, **P < 0.01.

In response to stimulation with CO₂, we determined that clear odor-evoked changes in GCaMP6s florescence intensity could be observed from Gr1+ OSNs in WT and yellow mutants (Fig. 3b). Strikingly, maximum odor-evoked changes in fluorescence from these cells were much higher in yellow mutants relative to WT (Fig. 3c). We next quantified maximum changes in GCaMP6s fluorescence intensity in response to stimulation with a range in CO₂ concentrations reflective of those found in diluted human breath (0, 0.1, 0.5, and 1% CO₂). We found that GCaMP6s responses were significantly elevated from Gr1+ OSNs in yellow mutants (Fig. 3d). In contrast, WT mosquitoes failed to exhibit major increases in GCaMP6s fluorescence across these concentrations (Fig. 3d). Notably, yellow mutant mosquitoes exhibited a significant, 2–4-fold increase in the maximum change in fluorescence above baseline (ΔF/F₀ values) relative to WT for each CO₂ concentration tested.

Introgression of QF2/QUAS transgenes into the yellow mutant A. aegypti strain thus enhances the sensitivity of transcuticular imaging of CO₂-evoked neural activity in this mosquito species over a range of gas concentrations.

Excision of 3xP3 fluorescent markers resolves spurious labeling in the central mosquito brain

We next evaluated central projection patterns in these 3xP3-marked chemoreceptor-QF2 driver lines when crossed to our 15xQUAS-mCD8::GFP responder described above. We expected that mCD8::GFP labeling would be restricted to axonal projections from each OSN class. Strikingly, immunohistochemical analysis of orco^QF2–3xP3 > 15xQUAS-mCD8::GFP, Ir8a^QF2–3xP3 > 15xQUAS-mCD8::GFP, and Gr1^QF2–3xP3 > 15xQUAS-mCD8::GFP adult female brains revealed spurious red and green fluorescence throughout the central brain (Fig. 4a–c). This was broadly evident in multiple cell types across the brain, particularly in orco^QF2–3xP3 > 15xQUAS-mCD8::GFP and Gr1^QF2–3xP3 > 15xQUAS-mCD8::GFP genotypes, suggesting potential background interference in the expected pattern of QF2/QUAS-based transactivation. As all of our T2A-QF2 insertions included downstream fluorescent marker cassettes containing the 3xP3 synthetic promoter (Berghammer et al. 1999), which is a multimerized binding site for the paired-box transcription factor Pax6 involved in glial and neuronal development (Quiring et al. 1994; Suzuki et al. 2016), we suspected such aberrant expression patterns may be due to promiscuous 3xP3 enhancer activity operating at these genomic loci.

Fig. 4.

Cre-loxP-mediated excision of 3xP3 marker cassettes from chemoreceptor T2A-QF2 driver lines facilitates clear visualization of OSN axonal projections into the A. aegypti antennal lobe. Spurious fluorescent labeling resulting from immunohistochemical analysis of adult female brains from a) orco^QF2–3xP3 > 15xQUAS-mCD8::GFP; b) Ir8a^QF2–3xP3 > 15xQUAS-mCD8::GFP; and c) Gr1^QF2–3xP3 > 15xQUAS-mCD8::GFP genotypes. Anti-BRP (magenta), anti-GFP (green). Arrows indicate putative glial cells from background 3xP3-DsRed2 marker expression outside of the optic lobes in the central brain. d) Schematic of crossing scheme to facilitate Cre-loxP-mediated excision of 3xP3 marker cassettes from chemoreceptor T2A-QF2^3xP3 driver lines with canonical and noncanonical HDR integrations to yield marker-free T2A-QF2^MF driver lines for central brain imaging. e) Schematic of resulting orco^QF2-MF, Ir8a^QF2-MF, and Gr1^QF2-MF driver integrations. Positioning of loxP sites indicated by arrows for a–e). f, g) Maximum intensity projections for f) orco^QF2-MF > 30xQUAS-mCD8::GFP and g) Ir8a^QF2-MF > 30xQUAS-mCD8::GFP genotypes to visualize orco+ and Ir8a+ projections. h) Single posterior z-slice for the Gr1^QF2-MF > 30xQUAS-mCD8::GFP genotype to visualize Gr1+ projections is shown. Arrows indicate the expression of the 3xP3-ECFP transgenesis marker for the QUAS-mCD8::GFP responder transgene in the outer optic lobes. Antennal lobes are encircled in white in a–c) and f–h). Confocal imaging settings were identical for all brains. SEZ, subesophageal zone. Scale bars: 50 µm. Images were taken at 20× magnification in a–c) and 10× magnification in f–h).

To abrogate this effect, we next excised the floxed 3xP3 fluorescent marker cassettes from these initial T2A-QF2^3xP3 strains by crossing these genotypes to a germline Cre recombinase strain (exu-Cre) that we engineered (Supplementary Table 7 in Supplementary File 1) to express Cre recombinase under the control of maternal germline promoter exuperantia (Li et al. 2017). We crossed males of each T2A-QF2^3xP3 driver line (Ir8a^QF2–3xP3, orco^QF2–3xP3, Gr1^QF2–3xP3) to exu-Cre females and screened F₁ progeny for loss of the 3xP3 fluorescent eye markers (Fig. 4d). In the case of the Gr1^QF2–3xP3 line, the duplicated 3xP3-DsRed2 marker due to the noncanonical HDR event was incompletely removed in F₁ progeny, so progeny still containing visible DsRed2 or ECFP markers were mated to their exu-Cre + siblings to ensure complete excision of all markers. Using this approach, we successfully generated marker-free driver strains (orco^QF2-MF, Ir8a^QF2-MF and Gr1^QF2-MF) which were devoid of all 3xP3 fluorescent markers (Supplementary Fig. 1 in Supplementary File 1), with only T2A-QF2 and a downstream loxP site remaining after excision at each locus (Fig. 4e).

We next generated a series of A. aegypti genotypes for imaging central brain neuroanatomy: orco^QF2-MF > 30xQUAS-mCD8::GFP, Ir8a^QF2-MF > 30xQUAS-mCD8::GFP, and Gr1^QF2-MF > 30xQUAS-mCD8::GFP. Each of these A. aegypti strains had 1 copy of each marker-free QF2 driver and 2 copies of the 15xQUAS-Syn21-mCD8::GFP-p10 transgene (i.e. for a cumulative dosage of 30xQUAS to express membrane-tethered GFP to label OSNs). Using immunohistochemistry analyses with a primary antibody directed against the presynaptic protein Bruchpilot (BRP) (Hofbauer et al. 2009) to demarcate glomerular boundaries of neuropils in the antennal lobe, and an anti-GFP antibody to amplify mCD8::GFP signal, we clearly labeled OSN axonal projections in the central A. aegypti brain without the spurious labeling seen with the 3xP3-marked driver lines. Of note, we determined orco+ neurons innervate a majority of antennal lobe glomeruli (Fig. 4f) and send axonal projections to the subesophageal zone (SEZ) (Supplementary Fig. 3 in Supplementary File 1). In contrast, Ir8a+ neurons project to several glomeruli in the posterior–lateral region of the antennal lobe (Fig. 4g), while Gr1+ neurons innervate a single glomerulus positioned deep in the posterior of the A. aegypti antennal lobe (Fig. 4h). These GFP labeling patterns were specific to each marker-free driver line, and we did not observe any apparent background fluorescence derived from OSNs innervating the peripheral sensory organs and antennal lobes of our 30xQUAS-mCD8::GFP only control strain (Supplementary Fig. 2a–d in Supplementary File 1).

We conclude that 3xP3 markers that are commonly used for A. aegypti transgenesis have the potential to induce spurious background expression in the central brain and that this can be resolved by their precise excision. We demonstrate that application of this method facilitates cell-type-specific labeling of OSN projections in the A. aegypti central brain with clarity.

An in vitro receptor-to-glomerulus map as the basis for improved A. aegypti antennal lobe annotation

We next performed confocal imaging and complete 3D morphological reconstructions of antennal lobes on replicate brain samples with labeled projections from orco+, Ir8a+, and Gr1+ OSNs using our marker-free T2A-QF2 A. aegypti genotypes. Across this dataset, we defined ∼79 total glomeruli (79 ± 3, mean ± SEM) in each reconstructed antennal lobe (Supplementary Fig. 4a in Supplementary File 1). This count was consistent with our previous estimate of ∼80 total glomeruli constituting the female A. aegypti antennal lobe based on reconstructions with synaptic staining alone (Shankar and McMeniman 2020). On average, we counted ∼63 orco+ glomeruli (63 ± 1, mean ± SEM), 15 Ir8a+ glomeruli, and 1 Gr1+ glomerulus in the reconstructed antennal lobes from each genotype (Supplementary Fig. 4b in Supplementary File 1).

Using a systematic reference key for A. aegypti antennal lobe nomenclature (Shankar and McMeniman 2020), we previously determined that 63 out of 80 total glomeruli (∼79%) could be found in stereotypical spatial positions in this mosquito species based on synaptic staining with anti-BRP antibody (nc82) alone. We determined here with transgenic labeling that cross-referencing BRP and GFP signals markedly improved our ability to define glomerular boundaries and discern the spatial arrangement of glomeruli within the antennal lobe in vitro. Indeed, we determined that 74 out of the 79 total glomeruli (∼94%) that we annotated could now be assigned to spatially conserved positions across reconstructions (Supplementary Fig. 4c and d in Supplementary File 1). A small proportion of glomeruli in each antennal lobe could not be reliably assigned to conserved spatial positions using our reference key and were classified as variant glomeruli (Supplementary Fig. 4c and e in Supplementary File 1).

We also determined that the region of the antennal lobe previously classified as the Johnston's Organ Center (JOC) (Ignell et al. 2005) appears to comprise of multiple discrete glomeruli in the postero-ventral and ventro-central groups of our updated atlas (Shankar and McMeniman 2020) that are innervated predominantly by orco+ neurons and to a lesser extent Ir8a+ neurons (Supplementary Fig. 4c in Supplementary File 1). Furthermore, volumetric analysis of glomeruli revealed that the Gr1+ glomerulus, denoted in both our atlas and prior atlases as MD1 (Ignell et al. 2005; Shankar and McMeniman 2020) or Glomerulus 1 (Herre et al. 2022), is the largest glomerulus in the antennal lobe (Fig. 5). Using spatial mapping, we also estimate a subset of 6 out of these 74 spatially invariant glomeruli are putatively both orco+ and Ir8a+ (Fig. 5 and Supplementary Fig. 4c and d in Supplementary File 1), consistent with recent observations of co-expression of these 2 genes in some OSN types (Herre et al. 2022).

Fig. 5.

Gr1+ OSNs innervate the largest glomerulus in the A. aegypti antennal lobe. a) 3D reconstructed model of the left antennal lobe of a female A. aegypti mosquito as seen from the anterior, lateral, posterior and medial perspectives. Template genotype for model: orco^QF2-MF > 30xQUAS-mCD8::GFP. Landmark glomeruli are indicated. b) Glomerular volumes from the female left antennal lobe. Mean volumes ± SEM from spatially invariant glomeruli are plotted, n = 7 brains. Volumes varied significantly (1-way ANOVA, P < 0.0001). Tukey's multiple comparison test, P < 0.05 for all comparisons to the CO₂ receptor glomerulus MD1.

The refined receptor-to-glomerulus map presented here thus reveals OSN populations expressing these 3 divergent chemoreceptors project centrally to defined regions of the A. aegypti antennal lobe.

The largest glomerulus in the A. aegypti antennal lobe detects carbon dioxide

To test whether the MD1 glomerulus which receives innervations from Gr1+ OSNs responds to CO₂, we generated mosquitoes suitable for central calcium imaging from the antennal lobe. To do this, we crossed our Gr1^QF2-MF driver and 15xQUAS-GCaMP6s responder lines to make Gr1^QF2-MF > 30xQUAS-GCaMP6s. This strain has 1 copy of the Gr1^QF2-MF driver and 2 copies of the 15xQUAS-GCaMP6s transgene to express GCaMP6s (Chen et al. 2013) in the cytoplasm of Gr1+ OSNs. Head-tethered A. aegypti females of this genotype with surgically exposed antennal lobes bathed in saline were then stimulated with different concentrations of CO₂. Consistent with observations from peripheral calcium imaging indicating Gr1+ neurons are tuned to CO₂, MD1 exhibited strong increases in odor-evoked GCaMP6s activity in response to stimulation with this gas (Fig. 6a–c). We then quantified the maximum change in GCaMP6s fluorescence intensity in response to stimulation with a CO₂ concentration series of 0.1, 0.5, and 1% CO₂ and determined that MD1 exhibits dose-dependent responses at this level of olfactory coding (Fig. 6d).

Fig. 6.

Gr1+ OSNs that project to the antennal lobe respond to CO₂. a) 3D reconstruction of the A. aegypti left antennal lobe. Gr1+ neurons project to a large posteriorly positioned glomerulus called MD1. b) GCaMP6s fluorescence in the right MD1 glomerulus before (left panel) and during 1% CO₂ stimulation (right panel). c) GCaMP6s traces after a 1% CO₂ pulse for 1 s (red bar) in the right MD1 glomerulus (Gr1^QF2-MF > QUAS-GCaMP6s). The solid line represents the mean and the shaded area the standard error of the mean (n = 9). d) Maximum change in fluorescence in the right MD1 glomerulus in response to 1-s pulses of increasing CO₂ concentration. The dots represent the mean, and the error bars the standard error of the mean (n = 9), paired t-test, **P < 0.01. e) CaMPARI2 green and red fluorescence in the Gr1+ MD1 glomerulus after stimulation with CO₂ and synthetic air. Right panels are heatmaps of red fluorescence intensity. MD1 from the left antennal lobe was imaged at 63× magnification. Scale bars: 10 µm. f) CaMPARI2 photoconversion values in MD1, mean R/G values ± SEM plotted, n = 3–5 brains per stimulus, Mann–Whitney U test, *P = 0.037.

We also applied the calcium-modulated photoactivatable ratiometric indicator (CaMPARI2) (Moeyaert et al. 2018) to record CO₂ activity from this glomerulus. CaMPARI2 photoconverts from green to red when simultaneously exposed to 405-nm light and high levels of calcium (Fosque et al. 2015; Moeyaert et al. 2018) and has previously been applied for activity-dependent neural labeling only in other model organisms such as mice, zebrafish, and flies. We generated a 15xQUAS-CaMPARI2 line via Mos1 mariner transposition (Supplementary Table 7 in Supplementary File 1) and then made a genotype for CaMPARI2 imaging with 1 copy of the Gr1^QF2-MF driver 2 copies of this responder transgene (Gr1^QF-MF > 30xQUAS-CaMPARI2) to express CaMPARI2 in the cytoplasm of Gr1+ OSNs. Consistent with GCaMP6s imaging results, MD1 exhibited a significantly higher rate of CaMPARI2 photoconversion in CO₂-stimulated mosquitoes vs those that were stimulated with synthetic air (Fig. 6e and f), validating the CO₂ sensitivity of this glomerulus.

These optimized genetic tools therefore facilitate clear imaging of odor-evoked activity from the A. aegypti antennal lobe and illustrate that the CO₂ receptor glomerulus MD1 which receives innervations from Gr1+ OSNs is sensitively tuned to carbon dioxide.

Discussion

The yellow fever mosquito A. aegypti is an emerging model system for invertebrate neurobiology. There has been a proliferation of sensory biology studies in this disease vector over the past decade (DeGennaro et al. 2013; McBride et al. 2014; McMeniman et al. 2014; van Breugel et al. 2015; Duvall et al. 2019; Raji et al. 2019; Jové et al. 2020; Melo et al. 2020; De Obaldia et al. 2022; Herre et al. 2022; San Alberto et al. 2022; Sorrells et al. 2022; Zhao et al. 2022; Chandel et al. 2024; Rouyar et al. 2024). To complement these efforts, here we report optimized genetic tools for neuroanatomical and functional mapping of the A. aegypti olfactory system. CRISPR–Cas9-mediated HDR has been an extremely important tool to generate T2A-QF2 in-frame fusions, rapidly expanding the number of cell-type-specific driver lines available in this mosquito species (Matthews et al. 2019; Jové et al. 2020; Zhao et al. 2021, 2022; Herre et al. 2022; Laursen et al. 2023). However, our study revealed several important issues for consideration when performing these types of transgenic manipulations in A. aegypti.

Firstly, we determined that when using circular donor template for CRISPR–Cas9 HDR, this can result in donor cassette integration either via canonical HDR, or alternatively via noncanonical HDR involving duplication of the intended T2A-QF2 integration cassette separated by intervening donor plasmid sequence. While both integration events yield precise T2A-QF2 in-frame fusions, duplication of the donor cassette and insertion of intervening vector backbone into the integration site may complicate molecular genotyping and cause other undesired effects. Including a second fluorescent marker in the plasmid backbone outside of the homology arms enabled us to detect this mode of integration. During gene targeting, we only observed noncanonical HDR events for Gr1, a single canonical HDR event for orco, and a mixture these HDR repair outcomes for Ir8a. While it is evident from this study that these differing types of HDR repair outcome are possible across these select target genes, additional studies are required to discern the relative frequency of canonical vs noncanonical HDR when using circular donor template at an expanded number of genetic loci in this mosquito.

We further demonstrated that strategic positioning of loxP sites surrounding the 3xP3 marker in the intended integration cassette can further be used to remove these florescent markers, including the second T2A-QF2 copy, and any extraneous vector backbone sequence integrated during multiplexed noncanonical HDR events. Evidence of multiple insertions of donor cassettes and plasmid backbone has been also observed in Drosophila and human cells (Bier et al. 2018; Bandyopadhyay et al. 2021; Zirin et al. 2022; Loehlin et al. 2023), suggesting that this mode of integration may be a common phenomenon when using circular donor template for CRISPR–Cas9-mediated HDR. We therefore highly recommend usage of the above design features where possible when employing circular HDR donor constructs for use with CRISPR–Cas9-mediated HDR in A. aegypti or other species.

We initially applied our transgenic reagents for functional imaging from the peripheral nervous system and observed that strong melanization of the A. aegypti cuticle dampened CO₂-evoked GCaMP6s responses in WT females. Introgression of our lines into a yellow mutant background (Li et al. 2017) allowed for a 2–4-fold increase in the signal recorded from CO₂-sensitive neurons in the maxillary palps and a larger dynamic range of responses along a CO₂ dose–response curve. This optimized method will thus facilitate future calcium imaging of odor-evoked activity with other subsets of A. aegypti OSNs located on maxillary palps or other sensory appendages in responses to stimulation with varied odorants, including components of the human volatilome (Rankin-Turner and McMeniman 2022). This technique may also facilitate high-throughput screens to determine the ligand tuning dynamics of different subsets of OSNs. Calcium imaging can be carried out in multiple sensilla housing various OSN classes simultaneously within a field of view, as opposed to single sensillum recordings (Afify and Potter 2022). Additionally, the increased signal obtained in yellow mutants during calcium imaging recordings will facilitate identification of neurons that show low responses to certain ligands that may be missed if imaging was carried out in a WT background.

Introgression of other driver and responder transgenes into the A. aegypti yellow mutant background (Li et al. 2017), with its lighter cuticular pigmentation, may increase the accuracy of peripheral cell counts particularly for sparser driver lines, as well as facilitating clear visualization of OSNs during antibody labeling and RNA in situ hybridization (Herre 2023a,b). Finally, application of pigmentation deficient yellow mutants has recently been employed in A. coluzzii to improve optical accessibility of the cuticle to visualize salivary gland colonization by Plasmodium berghei sporozoites (Klug et al. 2022), indicating the broad applicability of this approach to facilitate studies in both vector biology and neuroscience. However, as the yellow gene has been implicated in other insects in various physiological functions including cuticular pigmentation via production of dopamine melanin (Geyer et al. 1986; Wittkopp et al. 2002), binding dopamine or other biogenic amines (Xu et al. 2011), and egg chorion morphology (Noh et al. 2021), these potential factors should be considered by researchers when using yellow mutants for experimentation.

During evaluation of these transgenic reagents for central brain imaging, we determined that the 3xP3 markers typically used in insect transgenesis (Berghammer et al. 1999) induced spurious background fluorescence in the A. aegypti central brain when integrated as part of the T2A-QF2 in-frame fusion cassette at these 3 target genomic loci. Of note, removing these 3xP3 markers by Cre-loxP-mediated cassette excision (Häcker et al. 2017) to generate marker-free T2A-QF2 driver lines solved this problem. As such, given the widespread use of 3xP3 fluorescent markers for neurogenetic tools in A. aegypti (Matthews et al. 2019; Jové et al. 2020; Zhao et al. 2021, 2022; Herre et al. 2022; Laursen et al. 2023), we suggest that future designs of T2A-QF2 in-frame fusions in this mosquito species incorporate marker transgenes flanked by loxP sites to facilitate the flexibility to remove them for central neuroanatomical and functional imaging studies. This may be especially important within the context neurogenetic studies that use calcium imaging, optogenetics, and neuronal silencing in the A. aegypti nervous system where cell-type specificity of transgene expression and low background signal is crucial.

Using our optimized transgenic tools and immunohistochemistry, we could reliably identify 74 glomeruli in the female A. aegypti antennal lobe that were spatially conserved in our in vitro preparations by cross-referencing nc82 and GFP channels to demarcate glomerular boundaries. We determined that Ir8a+ OSNs innervate 15 glomeruli mostly localized to the posterolateral antennal lobe region, while Gr1+ OSNs innervate a single mediodorsal glomerulus called MD1 (Shankar and McMeniman 2020). We also showed that orco+ OSNs innervate a majority of spatially invariant antennal lobe glomeruli (56/74) that we identified and additionally project to the taste center of the mosquito brain known as the SEZ. A similar innervation pattern of orco+ OSNs to the SEZ from labellar neurons was observed in A. coluzzii (Riabinina et al. 2016), where it was hypothesized that this brain region may serve as center for gustatory and olfactory integration. Based on our observations of peripheral innervation of the A. aegypti labella by orco+ OSNs, future anterograde tracing is required to discern if this peripheral population of labellar neurons also project to the SEZ in this mosquito. Furthermore, whether the SEZ similarly integrates smell and taste information in A. aegypti remains unexplored. Finally, we estimate that a small subset of glomeruli in the A. aegypti antennal lobe co-express Ir8a and orco based on overlapping patterns of glomerular labeling in this brain center, as antennal immunostaining experiments with complementary T2A-QF2 driver reagents developed by others previously suggested (Herre et al. 2022).

Reduced or more variable numbers of antennal lobe glomeruli and receptor-to-glomerulus labeling patterns using T2A-QF2 in-frame fusions in A. aegypti have been recently described by 2 complementary studies, which found ∼60 (Zhao et al. 2022) or 60–72 total glomeruli (Herre et al. 2022). The upper range of these estimates is concordant with the 74 spatially invariant glomeruli that we observed in this study. We find that the use of our marker-free T2A-QF2 driver strains devoid of background interference from 3xP3 fluorescent markers, and a cumulative 30 copies of membrane-tethered GFP with translational enhancers (Pfeiffer et al. 2012) to strongly label OSNs, facilitates a clearer delineation of glomerular boundaries within the A. aegypti antennal lobe. As the repertoire of cell-type-specific driver lines for A. aegypti chemoreceptors increases, the annotation of glomeruli detailed here may be further refined. Indeed, the design principles described here to generate marker-free T2A-QF2 drivers devoid of background interference from transgenesis markers will be useful toward that end.

Moving forward, systematic receptor-to-glomerulus mapping in A. aegypti will help to discern whether certain glomeruli adjacent to one another represent different glomeruli or alternatively fragmented arborizations of the same glomerulus. Such morphological distortions may explain differences in the number of glomeruli observed across in vitro studies. Since additional improvements toward understanding the stereotypy of this brain region will lead to changes in glomerular numbering and naming over time as observed iteratively for Drosophila melanogaster (Stocker et al. 1990; Laissue et al. 1999; Couto et al. 2005; Fishilevich and Vosshall 2005; Tanaka et al. 2012; Grabe et al. 2015; Task et al. 2022), we have included raw confocal stacks and annotation files (Supplementary data) to facilitate future supplementary analysis or reannotation by the community.

We confirmed that the CO₂ receptor glomerulus MD1 is the largest neuropil in the A. aegypti antennal lobe (Ignell et al. 2005; Shankar and McMeniman 2020). The large volume of MD1 may indicate extensive synaptic connectivity between CO₂ receptor neurons, local interneurons, and projection neurons innervating higher-order olfactory processing centers of the mosquito brain. This likely reflects the critical importance of CO₂ to multiple facets of A. aegypti host-seeking behavior (Carlson et al. 1973; Geier et al. 1999; Bernier et al. 2003; Dekker et al. 2005; Dekker and Cardé 2011; McMeniman et al. 2014; Vinauger et al. 2019, #33; Sorrells et al. 2022; Chandel et al. 2024). We also reported for the first time the use of the calcium-modulated photoactivatable ratiometric indicator (CaMPARI2) (Moeyaert et al. 2018) in a nonconventional model organism. While CaMPARI2 cannot be used to record real-time neuronal activity like GCaMP indicators, it still provides a useful tool to study the activation of glomeruli deep in the antennal lobe where live imaging with GCaMP may be challenging and strongly affected by movement artifacts. Alternatively, this sensor may have utility for activity-dependent labeling in other brain regions in conjunction with other cell-type-specific or pan-neuronal drivers developed for A. aegypti (Matthews et al. 2019; Jové et al. 2020; Zhao et al. 2021, 2022; Herre et al. 2022; Laursen et al. 2023).

In this study, we generated disruptive T2A-QF2 in-frame fusions in coding exons of orco, Ir8a, and Gr1 and demonstrated the utility of these drivers when used as heterozygotes for functional imaging and neuroanatomy. However, it is worth noting that when using these drivers as homozygotes that loss-of-function phenotypes in these olfactory genes may be observed. When expanding this method to additional target loci, such as those implicated in development or physiology, careful consideration of whether the target gene is haplosufficient for the phenotype of interest is required. In such cases, T2A-QF2 in-frame fusions can be directed toward to the C terminus of the endogenous coding region in the target gene to reduce chances of gene disruption (Zhao et al. 2021; Herre et al. 2022).

The optimized genetic tools we report here may therefore be highly useful to study the molecular and cellular basis of A. aegypti olfaction including identification of chemosensory circuitry that mediates attraction to humans, oviposition sites, and mates. High-resolution studies of mosquito olfactory biology using neurogenetics may reveal rational targets for development of interventions to combat mosquito-borne diseases transmitted by A. aegypti such as dengue, Zika, and chikungunya.

Data availability

Strains and plasmids are available upon request. The authors affirm that all data necessary for confirming the conclusions of the article are present within the article, figures, and supplementary material. All raw data supporting these analyses have been deposited publicly in the Johns Hopkins Research Data Repository at: https://doi.org/10.7281/T1/HSROGO.

The full sequences of plasmids generated in this study have been deposited to GenBank: pBB-AaGr1 (accession no. PQ659180), pBB-AaOrco (accession no. PQ671723), pBB-AaIR8a (accession no. PQ671724), pBB (accession no. PQ671725), pMos-loxP-ECFP-loxP-15xQUAS-CaMPARI2 (accession no. PQ671726), pMosECFP-15xQUAS-GCaMP6s (accession no. PQ671727), pMosECFP-15xQUAS-mCD8GFP (accession no. PQ671728), and pMosEYFP-exu-Cre (accession no. PQ671729).

Supplemental material available at G3 online.

Acknowledgments

The authors thank N. Kizito, B. Natarajan, W. Okoth, H. Rosado, G. Nasir, M. Gebhardt, V. Balta, A. Ellison, and B. Burgunder for expert technical assistance; R. Harrell (UM-ITF) for mosquito embryonic microinjection services; S. Seo and A. Hammond for help with transgene mapping; C. Potter and E. Schreiter for constructs; and C. Huang, M. Schnitzer, C. Dan, and V. Jayaraman for guidance on surgical preparations. The mosquito template in Fig. 2 was created for us by Biorender.com.

Funding

This research was supported by funding from the National Institutes of Health (R21AI139358 and R21AI146450), United States Agency for International Development (AID-OAA-F-16-00061) and Centers for Disease Control and Prevention (200-2017-93143) to CJM. GMT and MPW were supported by postdoctoral fellowships from the National Institutes of Health (T32A1007417). The authors acknowledge generous support to CJM from Johns Hopkins Malaria Research Institute (JHMRI) and Bloomberg Philanthropies. SS, GMT, and DG were supported by JHMRI Postdoctoral Fellowships. OSA and ML were supported in part by awards from the National Institutes of Health (R01AI151004, R01AI148300, and R01AI175152) awarded to OSA. Microscopy infrastructure at Johns Hopkins School of Medicine Microscope Core Facility used in this research was supported by the National Institutes of Health NCRR (S10OD016374 and S10OD023548). The authors thank T. Shelley at for fabrication services supported by a NINDS Center grant from the National Institutes of Health (NS050274).

Author contributions

Conceptualization: SS, DG, GMT, and CJM; data curation, formal analysis, methodology, visualization, and writing—original draft: SS, DG, GMT, MPW, and CJM; funding acquisition: OSA and CJM; investigation: SS, DG, GMT, EDS, MPW, and CJM; project administration: CJM; resources: ML and OSA; and writing—review and editing: all authors.

Author notes

Shruti Shankar and Diego Giraldo contributed equally.

Conor J. McMeniman Lead contact.

Conflicts of interest: OSA is a founder of Agragene, Inc. and Synvect, Inc. with equity interest. The terms of this arrangement have been reviewed and approved by the University of California, San Diego, in accordance with its conflict of interest policies. All other authors declare no conflicts of interest.