Abstract
Background
Deciphering how developmental gene-regulatory programs interface with lineage-specific reproductive novelties in Bemisia tabaci demands a temporally resolved, integrative molecular framework. Using deep miRNA sequencing, high-throughput transcriptomics, quantitative proteomics, two-dimensional proteo-mapping, and spatially multiplexed FISH, we delineate the circuitry governing whitefly ontogeny.
Results
This atlas uncovers a previously unrecognized morphological innovation: a transporter-enriched egg stalk operating as an active, metabolically competent nutrient-harvesting appendage. Integrated temporal profiling indicates that the strongest molecular reprogramming in B. tabaci occurs at hatching and adult emergence, consistent with conserved developmental transition points described in other insects. Conserved miRNAs (Btab-mir-34 and Btab-mir-2944b) were associated with early developmental transitions, whereas clade-restricted miRNAs (Btab-mir-307a, Btab-mir-352a, and Btab-mir-107a) were predicted to regulate metabolic, detoxification, and chemosensory networks. Spatial FISH supports that Btab-novel-mir-018a represses vitellogenin during late-nymphal stages, establishing a developmentally gated post-transcriptional module relieved at adult emergence. Comparison of egg versus isolated stalk tissue proteomics revealed a transporter-enriched molecular signature for the B. tabaci egg stalk, extending earlier physiological evidence for pedicel-mediated water and solute uptake. Localization of NaPi-III and RNAi-associated egg viability phenotypes further supports a transport-related role for this structure during embryogenesis.
Conclusion
Collectively, this study provides a stage-wise molecular reconfiguration framework toward understanding the developmental and evolutionary architecture of B. tabaci. Furthermore, preliminary evidence for egg-stalk-associated transport identifies a candidate ontogeny-specific process that may inform future precision pest-management strategies.
Supplementary Information
The online version contains supplementary material available at https://doi.org/10.1186/s12864-026-13031-z.
Keywords: Bemisia tabaci, MiRNA-seq, MRNA-seq, Proteomics, Stage-resolved ontogeny, Egg stalk, And FISH
Background
The sweet potato whitefly, Bemisia tabaci, is among the most deleterious pests affecting crops worldwide, attributable to its extensive host range, rapid population growth, insecticide resistance, and exceptional capacity to transmit plant viruses [1, 2]. Instead of a single species, B. tabaci is presently regarded as a cryptic species complex, with this concealed diversity contributing to variations in host utilization, invasion efficiency, resistance traits, and vector competence [3]. Genomic analyses further suggest that its ecological success is supported by molecular complexity associated with detoxification, host adaptation, and virus transmission [4]. Indeed, the success of B. tabaci depends on precisely regulated, stage-specific processes, including embryogenesis, nymphal development, molting, cuticle formation, metabolism, and maturation into adulthood. Distinct transcriptomic signatures observed across the egg, nymph, and adult stages suggested that these developmental transitions were governed by distinct transcriptional programs and their regulatory molecular mechanisms [5]. For example, functional investigations reveal that genes involved in the miRNA pathway, such as Dicer1 and Argonaute1, are vital for normal survival and fecundity, whereas Dnmt1 is crucial for egg viability and early embryonic development, emphasizing the importance of post-transcriptional and epigenetic regulation in whitefly ontogeny [6–8]. Although hormonal and miRNA-mediated developmental regulation are well established in model insects, these integrated regulatory networks remain inadequately understood in B. tabaci, thereby limiting our capacity to elucidate its developmental plasticity and identify stage-specific molecular targets for sustainable pest management [6, 9].
Previous investigations into B. tabaci have focused primarily on transcriptomics or proteomics in isolation, leading to the identification of stage-specific cuticle proteins, detoxification enzymes, and stress‒response factors [5]. However, these singular approaches fail to elucidate the hierarchical relationships between miRNAs and their mRNA targets, nor do they comprehensively capture developmental processes at the proteome level. While well-conserved miRNAs are recognized for serving as developmental checkpoints, the roles of novel, lineage-specific miRNAs in B. tabaci ecological specialization, particularly in detoxification and chemosensation, are not well defined [10, 11]. Moreover, in Tribolium castaneum, for example, transcriptome-wide profiling identified major reprogramming between late eggs and early larvae and again between late pupae and early adults, including coordinated changes in DNA replication, cell-cycle, and endocrine pathways [12]. More broadly, the juvenile-to-adult transition is governed by conserved regulatory logic across hemimetabolous and holometabolous insects [13].
In B. tabaci, previous physiological studies demonstrated that the egg pedicel mediates water uptake and can transport solutes into the developing egg [14, 15], but the molecular composition of this structure remains unresolved. The evolution of insect reproductive structures reflects a range of survival strategies developed for early life, resource acquisition, and ecological adaptation. In hemipterans, egg-associated features are often limited to small, passive pedicels or anchoring filaments; however, physiological studies indicate that some lineages utilize host resources during embryogenesis. For instance, whitefly eggs are hydrated by water drawn from plant tissue through the pedicel [14], suggesting lineage-specific survival mechanisms at this stage. In this context, the present study examines whether the B. tabaci egg stalk carries a distinct transporter-enriched molecular profile consistent with the previously proposed physiological role of the pedicel. Furthermore, the abundance of Na⁺/K⁺-ATPase and other solute transport-associated proteins is consistent with the specialized transport-related role of ion pumps in insect epithelia and resource mobilization [16]. The adaptation of these transporters to a reproductive appendage demonstrates the evolutionary repurposing of conserved osmoregulatory systems. Such gene repurposing in response to ecological pressures has parallels in other insects [17], supporting the notion that selection can rapidly redirect transporter functions to suit niche demands. In B. tabaci, stalk-mediated delivery of resources from the plant not only enhances survival but also facilitates prolific reproduction and rapid population spread. Provisioning the next generation with critical resources directly from the host plant increases hatching success and overall reproductive efficiency while buffering offspring against environmental variability. These benefits align with broader models in which resource allocation strategies influence fitness and life-history evolution [18], and are supported by evidence that disruption of key transporters, such as Na⁺/K⁺-ATPase, adversely affects insect reproductive output [19].
In this study, we combined small RNA sequencing, mRNA transcriptomics, and quantitative proteomics across all significant developmental stages of B. tabaci, namely, eggs, nymphal instars I–IV, puparia, and adults. This integrative approach provides a comprehensive molecular atlas highlighting that the most extensive transcriptomic, miRNA, and proteomic reconfiguration in B. tabaci occurs at hatching and adult emergence. The early phase is characterized by conserved miRNAs associated with genes involved in replication and biosynthesis, thereby maintaining embryonic stability. In contrast, the latter phase expressed novel miRNAs predicted to influence detoxification and sensory pathways critical for the ecological adaptation of adults. These findings illustrate that insect development is driven by a hierarchical regulatory cascade in which miRNAs orchestrate transcriptional programs that are accurately translated into the proteome. Our results help elucidate the molecular basis of insect development and identify regulatory targets with potential implications for managing whitefly populations. This study may open a new avenue for the development of a pest counter-mechanism that targets developmental biology rather than relying heavily on synthetic insecticide-based pest management.
Methods
Insect source
The B. tabaci collected from cotton (Gossypium hirsutum, variety Co17) fields at the Department of Cotton, Tamil Nadu Agricultural University (TNAU), Coimbatore, Tamil Nadu, India, was initially established in January 2022 in glasshouse facilities at the Department of Agricultural Entomology, TNAU, Coimbatore, Tamil Nadu, India. This population was maintained on potted cotton plants aged 35–50 d (Co17), providing optimal reproduction. This population was perpetually maintained in insect-proof rearing cages, with the old cotton plants replaced monthly with fresh samples under strictly controlled glasshouse conditions (30 ± 2 °C; 70 ± 5% humidity; 12 h light/dark cycle). Genomic DNA was extracted from representative samples for species identification and genetic lineage identification. This was followed by amplification of the mitochondrial cytochrome oxidase subunit I (mtCOI) gene using a universal primer set [20] by polymerase chain reaction (PCR) on a Mastercycler® X50 (Eppendorf, Germany). The resulting PCR amplicons were sequenced, and subsequent phylogenetic analysis confirmed the population’s affiliation with the Asia I genetic group, which aligned with established regional biotypes of B. tabaci.
Egg sampling and stalk dissection
Eggs of B. tabaci were carefully removed from infested host leaves using a microneedle with a tip diameter of approximately 0.22 µm (Techno, USA) under a stereomicroscope (Leica M205C, Leica Microsystems, Germany) and transferred directly to low-binding microcentrifuge tubes (Axygen, USA) containing chilled phosphate-buffered saline (pH 7.4; filter-sterilized, 0.22 µm; Sigma Aldrich, USA) and kept on ice during collection. For stalk collection, detached eggs were immediately transferred into sterile PBS and placed in a sterile cavity dish (Tarsons, India) maintained on ice to prevent desiccation. The attached stalks were then dissected under a microscope using a custom-made micro-knife fabricated from a sharpened tungsten wire, mounted on a micromanipulator. Each stalk was incised precisely at its basal junction without damaging the egg body, and the separated stalks were gently aspirated into chilled PBS using fine glass microcapillaries (Eppendorf, Germany). Dissected stalks were pooled in sterile, low-binding microcentrifuge tubes containing chilled PBS and kept on ice during collection. For proteomic analysis, samples were supplemented with a protease inhibitor cocktail (Sigma-Aldrich, USA) and stored at − 80 °C (Thermo Fisher Scientific, USA) until extraction.
Scanning electron microscope (SEM) analysis
Samples representing various developmental stages of B. tabaci were carefully prepared for SEM in accordance with established protocols. To remove surface contaminants, the samples were first rinsed with PBS, then gradually dehydrated using an ethanol series (30–100%; 10 min each). After dehydration, the samples were subjected to critical-point drying using a Quorum Technologies dryer (Quorum Technologies, UK) to maintain their fine surface features. They were then affixed to aluminum stubs with conductive carbon adhesive tabs to increase the electrical conductivity. To mitigate charging effects during imaging, the samples were sputter-coated with a thin (~ 10 nm) gold layer via a Quorum SC7620 sputter coater (Quorum Technologies, UK). An imaging was conducted using an SEM (ZEISS EVO MA 15, Carl Zeiss, Oberkochen, Germany) operating under high-vacuum conditions, with accelerating voltages ranging from 5 to 10 kV. A diverse array of low- and high-magnification micrographs was acquired to elucidate stage-specific surface characteristics, including cuticular sculpturing and setal distribution. Instrument parameters—such as accelerating voltage, working distance, and magnification—were carefully optimized for each sample to achieve optimal resolution while minimizing beam-induced damage.
Temporal multi-omics approaches
Insect sampling process
The B. tabaci used for omics studies were obtained from well-established laboratory colonies maintained in a glasshouse, ensuring genetic consistency and traceability throughout the research. The developmental stages of B. tabaci, including eggs (whole eggs, with stalk, and without stalk), nymphs (first to fourth instars), puparia, and adults, were collected directly from host plant foliage. Using a Leica stereomicroscope (Leica Microsystems GmbH, Wetzlar, Germany), individuals were carefully detached from the leaf surface with a fine 22-gauge needle to minimize physical damage. Immediately after collection, the insects were transferred into sterile, RNase-free microcentrifuge tubes prefilled with RNAlater® Stabilization Solution (Sigma‒Aldrich, USA). The samples were incubated at 4 °C overnight to allow thorough permeation of the preservative and then stored at − 80 °C in ultralow-temperature freezers (Thermo Fisher Scientific, USA) until further use. For small RNA studies, ~ 2000 eggs and ~ 100 individuals from each nymphal instar per replicate were processed independently to preserve microRNAs and other regulatory RNAs. For mRNA analyses, insects from each developmental stage were preserved under the same conditions, and total RNA was subsequently isolated to capture long transcripts for transcriptome profiling. All samples were replicated twice. For proteomic analyses, samples were collected as for RNA processing, stored in RNAlater®, briefly rinsed with 1% PBS after thawing to remove excess preservative, then homogenized under protease-inhibiting conditions, protein quantified, and analyzed by liquid chromatography-tandem mass spectrometry (LC–MS/MS). All the proteomics samples were replicated three times. This standardized workflow ensured reliable recovery of intact small RNAs, mRNAs, and proteins across all developmental stages of B. tabaci.
Small RNA sequencing
Total RNA enriched for miRNAs was isolated from B. tabaci using TRIzol® Reagent (Sigma-Aldrich, St. Louis, MO, USA) [21]. The small-RNA fraction was then recovered using the miRNeasy Micro Kit (Qiagen, Hilden, Germany). RNA purity was assessed spectrophotometrically with a NanoDrop 2000 (Thermo Fisher Scientific, Waltham, MA, USA), and RNA concentration was measured with the RNA Broad Range Assay Kit (Invitrogen, Thermo Fisher Scientific, Waltham, MA, USA) on a Qubit 4 Fluorometer (Thermo Fisher Scientific, Waltham, MA, USA). RNA quality was further verified by agarose gel electrophoresis (Bio-Rad, USA). Small-RNA libraries were prepared using the QIAseq miRNA Library Kit (Qiagen, Hilden, Germany). Library cleanup was carried out with QIAseq beads (Qiagen, Hilden, Germany), and fragment-size distributions were evaluated on an Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA) using High Sensitivity DNA chips (Agilent Technologies, Santa Clara, CA, USA). Sequencing was performed on an Illumina NovaSeq 6000 platform (Illumina, San Diego, CA, USA) to generate single-end reads for downstream miRNA analysis.
Transcriptomic analysis
Total RNA was extracted from developmental-stage B. tabaci samples stored in RNAlater®. RNA purity was assessed using a NanoDrop 2000 spectrophotometer, and RNA concentration was determined using the RNA Broad Range Assay Kit on a Qubit 4 Fluorometer. RNA integrity was evaluated by agarose gel electrophoresis and confirmed on an Agilent 2100 Bioanalyzer. The libraries were generated from high-quality RNA using the KAPA HyperPrep cDNA Synthesis Module (Roche Diagnostics, Basel, Switzerland), including poly(A) enrichment and cDNA library preparation. Libraries were purified using KAPA Pure Beads (Roche Diagnostics, Basel, Switzerland) and assessed on an Agilent 2100 Bioanalyzer with High Sensitivity DNA chips. Sequencing was carried out on an Illumina NovaSeq 6000 platform to generate paired-end transcriptomic reads.
Untargeted shotgun proteomic analysis
Protein extraction and digestion
Whitefly samples were homogenized in 10% trichloroacetic acid in acetone supplemented with 100 mM PMSF, 10 mM DTT, and a protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO, USA) [22]. Briefly, each preparation contained 500 ng tissue in 800 µL extraction solution. Protein precipitation was carried out at − 20 °C for at least 16 h, followed by centrifugation at 12,000 × g for 15 min. The pellet was washed three times with chilled acetone containing 10 mM DTT, air-dried, and dissolved in rehydration buffer. Extracted proteins were stored at − 80 °C until use. Protein concentration was determined using the Qubit Protein Assay Kit on a Qubit 4 Fluorometer. For mass spectrometry, 100 µg of protein was mixed with 50 µL RapiGest SF (Waters Corporation, Milford, MA, USA) and incubated at 65 °C for 30 min. Reduction was performed with 10 µL of 100 mM DTT for 30 min at 65 °C. Samples were then alkylated with 20 µL of 100 mM iodoacetamide for 30 min in the dark. Digestion was carried out with MS-grade RapiZyme Trypsin (Waters Corporation, Milford, MA, USA) at an enzyme-to-substrate ratio of 1:20 for 16 h at 37 °C in a dry bath (LABIOD®, Solan, Himachal Pradesh, India). Reactions were stopped by adding 1 µL formic acid. The resulting peptides were concentrated under vacuum using a vacuum concentrator (Eppendorf, Hamburg, Germany) and desalted on a C18 reverse-phase column (Waters Corporation, Milford, MA, USA).
LC–MS/MS analysis
Peptides were separated on a Waters UPLC system (Waters Corporation, Milford, MA, USA) using an Acquity UPLC BEH C18 column (75 µm × 150 mm, 1.7 µm; Waters Corporation, Milford, MA, USA). Mobile phase A consisted of LC–MS-grade water containing 0.1% formic acid, and mobile phase B consisted of acetonitrile containing 0.1% formic acid. The column temperature was maintained at 60 °C, and the gradient was programmed to increase phase B to 85%. The flow rate was set at 5 µL min⁻1 for the first 3 min, then increased to 200 µL min⁻1. Mass analysis was performed on a Xevo G2-XS mass spectrometer (Waters Corporation, Milford, MA, USA) equipped with an electrospray ionization source operating in positive mode. Instrument settings were as follows: low energy, 4 V; transfer, 4 V; cone voltage, 40 V; trap collision energy ramp, 15–40 V; scan time, 0.5 s; and TOF range, 50–2000 Da. Leucine enkephalin (Sigma-Aldrich, St. Louis, MO, USA) at 200 pg µL⁻1 was used for external calibration, with lock-mass correction applied every 30 s. Spectra were acquired and processed using MassLynx version 4.0 (Waters Corporation, Milford, MA, USA). The analysis included three biological replicates and two technical replicates.
Protein identification
Raw files were processed in Progenesis QI for Proteomics version 4.0 (Nonlinear Dynamics, Newcastle, UK). Retention-time alignment was performed before peak extraction, and peak-picking sensitivity was set to 5 ppm. Features detected between 5 and 80 min and within an m/z range of 100–1600 were retained. Peptides that could not be assigned unambiguously to a single protein were excluded from quantitative analysis. Protein annotation was performed against a custom UniProt-derived database containing entries from Hemiptera, Aleyrodidae, and B. tabaci. Search parameters included carbamidomethylation of cysteine as a fixed modification; oxidation of methionine and N-terminal pyroglutamate formation from glutamine as variable modifications; tryptic specificity with up to two missed cleavages; precursor mass tolerance of 20 ppm; at least two peptides per protein; a minimum of two fragment ions per peptide; and at least five fragment ions per protein. Protein identifications were filtered at a 5% false discovery rate (FDR). The raw proteomics data were deposited in the ProteomeXchange Consortium via jPOSTrepo under accession number PXD06799.
2D-PAGE
Isoelectric focusing (IEF) was performed for egg samples (with and without stalk) on immobilized pH gradient (IPG) strips (pH 3–10, 7 cm; Bio-Rad, California, USA) using a PROTEAN® i12™ IEF cell (Bio-Rad, USA). A total of 50 µg of protein per sample was mixed with 125 µl of rehydration buffer and passively incubated for 3 h, followed by active rehydration at 50 V for 10 h. IEF was then carried out at 4,000 V for two hours until a total of 50 kWh was reached. Strips were stored at − 80 °C prior to the second dimension. For equilibration, strips were incubated for 20 min in 5 ml of equilibration buffer containing 10 mM DTT (50 mM Tris, 6 M urea, 30% glycerol, 2% SDS), followed by a second 20 min incubation in the same buffer containing 135 mM iodoacetamide. SDS-PAGE was performed on 12% acrylamide gels (Bio-Rad, USA) at 95 V for 45 min. Protein spots were visualized via silver staining.
Protein identification by MALDI-TOF/mass spectrometry
A total of nine protein spots, consisting of five derived from stalked eggs and four from unstalked eggs, were carefully excised from preparative two-dimensional electrophoresis (2-DE) gels using a 1.5-mm picking pen (The Gel Company, San Francisco, CA). The gel plugs were subjected to a series of washes: initially with ultrapure water, then with a 1:1 (v/v) mixture of 100 mM ammonium bicarbonate (NH₄HCO₃) and acetonitrile (ACN) for 10 min, followed by a wash with 100% ACN for 5 min. After dehydration, the plugs were incubated with 100 ng of modified trypsin (RapiZyme Trypsin, MS Grade, Waters Corporation, Milford, Massachusetts, USA) in a solution of 40 mM NH₄HCO₃ containing 10% ACN for 30 min at 4 °C, then subjected to overnight digestion at 37 °C. Post-digestion, peptides were sequentially extracted with solutions containing 50% ACN and 2.5% formic acid (FA), and 90% ACN and 0.1% FA, then pooled with the digestion supernatant and vacuum-dried. The peptide mixture was desalted through C18 ZipTips (Millipore, Billerica, MA) and further purified via a two-step process employing ultra-performance liquid chromatography (UPLC) on an Acquity UPLC system (Waters Corp., USA) with a C18 column (1.7 µm, 2.1 × 100 mm). Elution was performed with a linear gradient from 5 to 95% ACN containing 0.1% FA over 20 min at a flow rate of 0.3 ml min⁻1. Fractions containing peptide peaks, detected at 214 nm, were pooled and concentrated for mass spectrometric analysis.
In preparation for MALDI-TOF/TOF mass spectrometry, purified peptides (0.5 µl) were combined with an equal volume of a matrix solution consisting of 10 mg ml⁻1 α-cyano-4-hydroxycinnamic acid in a solvent mixture of 50% acetonitrile (CAN), 0.1% trifluoroacetic acid (TFA), and 1 mM ammonium phosphate. This mixture was then spotted onto a stainless-steel 384-well target plate. Mass spectra were acquired using an AXIMA Performance MALDI-TOF/TOF mass spectrometer (Shimadzu Kratos Analytical, UK), operated with a 337 nm nitrogen laser at a pulse rate of 50 Hz and a pulse width of three nsec. The instrument was configured in positive-ion reflector mode, covering a mass-to-charge ratio (m/z) range of 900–4000 Da, with an acceleration voltage of 20 kV, a reflector voltage of 26 kV, and an extraction delay of 200 nsec. Each spectrum represents an average of approximately 1,500–2,000 laser shots. External calibration was performed using the Peptide Calibration Standard II (Bruker Daltonics), while internal calibration used trypsin autolysis peaks at m/z 1045.5642 and 2211.1046 Da. For tandem mass spectrometry (MS/MS), precursor ions were automatically selected based on a signal-to-noise (S/N) threshold of 10 and fragmented via post-source decay (PSD). Metastable ion suppression was enabled, with a reflector voltage set at 1.9 kV, and an average of 3,000 laser shots per spectrum was acquired. The spectra obtained from both MS and MS/MS analyses were examined using GPS Explorer v3.5 (Shimadzu/Applied Biosystems). Subsequently, the data were searched against the National Center for Biotechnology Information (NCBI) non-redundant (nr) database utilizing the Mascot algorithm (Matrix Science, Boston, MA).
Gene isolation, cloning, and bacterial transformation
Total RNA was extracted from approximately 2000 freshly harvested eggs using TRIzol reagent following the manufacturer’s protocol. To minimize genomic DNA contamination, we treated the RNA with RNase-free DNase I (Thermo Fisher Scientific, USA), quantified it using a NanoDrop spectrophotometer, and further evaluated its quality via agarose gel electrophoresis. For the synthesis of first-strand complementary DNA (cDNA), one µg of DNase-treated RNA was used as the template, and a high-fidelity reverse transcriptase (PrimeScript, Takara, Japan) was used according to the manufacturer’s instructions. The resultant cDNA served as a substrate for amplification of two distinct transporter genes: the first, a sodium-dependent potassium transporter (approximately 1.6 kb); and the second, an uncharacterized protein homologous to potassium transporters (approximately 1.4 kb). Gene-specific primers were meticulously designed to amplify the entire coding sequences derived from the B. tabaci whole-genome datasets.
The amplification reactions were carried out using iProof™ High-Fidelity DNA Polymerase (Bio-Rad, USA) within 50 µl reaction volumes, containing 10–20 ng of cDNA template, 0.5 µM of each primer, 200 µM dNTPs, and 1X HF buffer. The thermal cycling parameters comprised an initial denaturation at 98 °C for 30 s, followed by 35 cycles of 98 °C for 10 s, 57 °C for 30 s, and 72 °C for 1.5 min, concluding with a final extension at 72 °C for 5 min. PCR products were analyzed by 1% agarose gel electrophoresis, with single discrete bands corresponding to expected amplicon sizes excised and purified using the NucleoSpin Gel and PCR Clean-up kit (Takara, Japan). Subsequently, the purified PCR products were cloned into pJET1.2 using the CloneJET PCR Cloning Kit (Thermo Fisher Scientific, USA) following the manufacturer’s instructions and were transformed into E. coli DH5α cells. The positive colonies were screened for recombinant inserts by colony PCR using vector-specific primers and further validated by plasmid extraction using the NucleoSpin® Plasmid Kit (Takara, Japan), followed by diagnostic restriction enzyme digestion and Sanger sequencing.
Heterologous expression and protein purification
Following the verification of the sequence, the inserts corresponding to the sodium-dependent potassium transporter were subcloned into the pET-28a(+) and subsequently introduced into E. coli BL21 (DE3). To confirm the presence of correct colonies, colony PCR and diagnostic restriction digestion were performed to verify the correct insert orientation and reading frame. For protein expression, single verified colonies were inoculated into LB broth containing kanamycin and induced with 0.1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) after an optical density (OD₆₀₀) of approximately 0.6 was reached. The cultures were later harvested by centrifugation, and the resulting pellets were resuspended in a lysis buffer composed of 50 mM Tris–HCl (pH 8.0), 300 mM NaCl, and 10 mM imidazole. Proteins were analyzed by SDS–PAGE to verify their purity and molecular weight. Finally, the purified fractions were quantified spectrophotometrically and stored at −80 °C for subsequent functional and structural analyses.
Stem‒loop quantitative PCR (qPCR)
Candidate miRNAs with strong stage-specific expression were selected for validation by stem–loop qPCR. Total RNA was extracted from three biological replicates using TRIzol® Reagent following the manufacturer’s instructions. Stem–loop primers were designed manually according to Chen et al. (2005) (Supplementary File 1). Reverse transcription was performed with the PrimeScript RT Reagent Kit (Takara Bio Inc., Kusatsu, Shiga, Japan) using 1000 ng total RNA in a 20 µL reaction containing miRNA-specific stem–loop RT primers. Quantitative PCR was carried out in 20 µL reactions using TB Green® Premix Ex Taq™ II (Takara Bio Inc., Kusatsu, Shiga, Japan) on a CFX Opus 96 Real-Time PCR System (Bio-Rad Laboratories India Pvt. Ltd.). Cycling conditions consisted of 95 °C for 2 min, followed by 40 cycles of 95 °C for 15 s, 57 °C for 15 s, and 72 °C for 25 s. A no-template control containing nuclease-free water was included in each run. Melt-curve analysis was performed using CFX Maestro software (Bio-Rad, USA) to confirm amplification specificity.
Whole-mount fluorescence in situ hybridization (FISH)
The procedure began with the fixation of samples, including eggs, instar IV nymphs, and adults. These samples were immersed in a 4% paraformaldehyde solution (CDH, New Delhi, India) prepared in 1 × PBS at room temperature for one h to preserve their structure. Probe sequences and final working hybridization concentrations are provided in Supplementary File 1. Following fixation, the samples were rinsed three times in 1 × PBS for five min each to remove any excess fixative. To permeabilize the tissue, proteinase K (20 mg/mL, Invitrogen, USA) was applied at a concentration between 20 µg mL⁻1 for 15 min at 37 °C, allowing the enzyme to break down cellular barriers. After permeabilization, the samples were refixed with fresh 4% paraformaldehyde solution for 10 min. Hybridization was performed by incubating samples overnight at 37 °C with four probes labeled with FAM and Cy5 dyes (BioServe, Hyderabad, India), targeting vitellogenin mRNA and NaPi-III transcripts with FAM, and Btab-novel-miR-018a and LOC109039444 with Cy5; probe sequences are in Supplementary File 1. The final hybridization concentration was 125 nM for each probe. Samples were pre-equilibrated in wash buffer containing 10% formamide in 2 × SSC and hybridized in buffer containg 10% dextran sulfate and 10% formamide in 2 × SSC to enhance probe stringency and signal specificity. To remove unbound probes, samples were washed twice in 10% formamide in 2 × SSC at 37 °C for 30 min, then rinsed in 2 × SSC. Sense-probe and no-probe controls were processed similarly to assess nonspecific fluorescence [23]. For mounting, a medium consisting of 90% glycerol and 10% PBS (1%) was freshly prepared. A small volume (20–40 µl) of mounting medium was placed between the spacers, and the samples were carefully transferred onto the microscopic slide under a stereo zoom microscope (Leica, Wetzlar, Germany) via a sterile fine 50 µm titanium needle (TED Pella, CA, USA), with the region of interest oriented toward the coverslip. The coverslip was gently lowered at an angle to avoid trapping air bubbles, keeping the sample submerged without flattening. The edges of the coverslip were sealed with clear nail polish. To protect fluorescence, the slides were stored flat in the dark at 4 °C and imaged within 24–48 h. Imaging was conducted on a Leica confocal microscope (Wetzlar, Germany) with sequential scanning: first, the FAM channel (green excitation and emission) was captured, then the Cy5 channel (far-red) was captured, with minimal laser power and exposure times. Complete z-stacks were acquired to capture signals at multiple depths.
RNAi in NaPi-III annotation
B. tabaci instar IV nymphs were enclosed in clip cages, and emergent adult individuals were collected at dawn. These pre-mated B. tabaci adults were subsequently subjected to an eight-hour feeding regimen on a diet laced with dsRNA. Thereafter, treated individuals were transferred onto untainted foliage within controlled enclosures to facilitate oviposition. Target region delineation, oligonucleotide design, amplification, and molecular cloning were consolidated into a streamlined workflow optimized for both in vitro and microbial dsRNA biosynthesis to generate dsNaPi-III and dsGFP constructs. A discrete NaPi-III locus unique to B. tabaci was identified using siRNA Wizard 3.1, and sequence specificity was verified by BLAST analysis against available vertebrate, invertebrate, and plant databases. For dsRNA biosynthesis, gene-specific primers were synthesized with a 5′ T7 promoter sequence and suitable restriction endonuclease sites for oriented ligation. In parallel, an irrelevant GFP fragment was amplified under the same strategy to generate dsGFP as a non-target negative control for feeding assays. Primer sequence information for both NaPi-III and GFP constructs is provided in Supplementary File 1. The resulting PCR products were purified, verified by agarose gel electrophoresis, and used as templates for downstream dsRNA production. Total RNA from approximately fifty adult B. tabaci specimens was extracted employing TRIzol reagent, and one microgram was reverse transcribed via the PrimeScript First-Strand cDNA Synthesis Kit (Takara, Japan). PCR amplification used 0.4 μM each modified primer, 0.26 mM dNTPs, 2 × HF buffer, and 2 U iProof HF polymerase in a 25 μL mixture, with cycling parameters of: 98 °C for 3 min; 35 cycles of 98 °C for 10 s, 60 °C for 30 s, 72 °C for 30 s; followed by a 10 min extension at 72 °C. The resultant NaPi fragment was excised from a 2% agarose gel using the NucleoSpin Gel and PCR Clean-up Kit (Takara, Japan). For microbial dsRNA generation, the PCR product was restricted with NotI and NcoI, directionally inserted into a dual-T7 expression vector, and verified via colony PCR and sequencing in E. coli DH5α. Recombinant constructs were subcloned into RNase III–deficient HT115(DE3) cells for IPTG-stimulated dsRNA expression. Induced cultures were harvested, lysed with TRIzol® and analyzed via agarose gel electrophoresis, while eggs laid after dsNaPi-III treatment and the corresponding controls were collected in parallel for total RNA extraction, cDNA synthesis using the same PrimeScript protocol, and subsequent qPCR (CFX96, Bio-Rad, USA) analysis of NaPi-III transcript abundance.
Bioinformatics analyses
Raw sequencing data from small RNA and mRNA libraries were processed using standard pipelines with meticulous version control to ensure reproducibility. For both the mRNA and small RNA datasets, adapter sequences were removed, and quality filtering was performed with Cutadapt v4.4 [24], preserving reads of at least 36 nucleotides for mRNAs and 17–30 nucleotides for small RNAs. The quality of the libraries was evaluated with FastQC v0.11.9 [25] and summarized through MultiQC v1.14 [26]. Transcriptome reads were quantified against the reference genome using STAR [27], and transcript-level abundances were aggregated to the gene level using the RSEM package. For miRNA sequencing, both known and novel candidates were identified using miRDeep2 v2.0.1.2 [28], with reference to the most recent miRBase release 22.1 [29]. The novel miRNAs were retained only when the canonical hairpin structures were predicted, the read abundance exceeded 10 reads per million, and detection was consistently observed across at least two biological replicates. Proteomic spectra were analyzed with amica software (v.3.0.1) [30], which conducted peptide-spectrum matching against the UniProt species-specific protein database, filtered identifications at a 1% false discovery rate via target–decoy methodologies, and normalized protein intensities. Protein intensity values were log2-transformed, and missing values were estimated through a downshifted normal distribution to simulate low-abundance signals.
Differential expression analysis
Differential expression analysis of the mRNA and miRNA datasets was conducted via DESeq2 v1.38.3 [31] within R software v4.5.1. Genes exhibiting low counts across replicates were filtered for analysis. The Wald test was applied for pairwise comparisons, whereas the likelihood ratio test (LRT) was used to evaluate global effects across developmental stages. Independent filtering and parametric dispersion estimation were employed. Significant differential expression was defined as Benjamini–Hochberg-adjusted p < 0.05 and an absolute log2FC ≥ 1. Proteomic differential abundance was derived from the statistical module within amica through empirical Bayes moderation. Statistical significance was similarly defined as adjusted p < 0.001 and |log2FC|≥ 1, depending on the dataset’s variance.
Integration of the miRNA‒mRNA‒proteome through inverse relationships
The miRNA–mRNA target interactions were predicted via a consensus approach, leveraging TargetScan v8.0 [32], miRanda v3.3a [33], and RNAhybrid v2.1.2 [34]. Predictions were retained only if supported by a minimum of two algorithms. For novel miRNAs, predictions were confined to high-confidence sites with ≤ − 20 kcal mol−1 binding energies. Correlation analyses were performed across matched samples via Spearman’s rank correlation in R. Putative inhibitory relationships were defined as having correlation coefficients ≤ − 0.4 and Benjamini–Hochberg adjusted p < 0.05. MiRNAs were classified into four functional categories based on the presence or absence of inversely regulated mRNA and protein targets, encompassing both known and novel candidates.
Functional annotation and pathway analyses
Functional enrichment analysis was performed using the clusterProfiler v4.6.2 [35]. Both the Gene Ontology (GO) [36] and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathways [37] were analyzed using hypergeometric models, with a false discovery rate (FDR) of < 0.05 deemed significant. Gene set enrichment analysis was conducted using fgsea v1.24.0 [38] to capture pathway-level alterations without relying on fold-change thresholds. Analyses were conducted separately for transcriptomic and proteomic datasets and jointly for miRNA–mRNA–protein triplets.
Network construction and stage-specific dynamics
Regulatory networks were constructed in Cytoscape v3.10.1 [39] with directed edges representing miRNA–mRNA and mRNA–protein interactions. Edge weights reflect combined evidence scores that include prediction confidence, differential expression statistical significance, and correlation strength. Stage-specific contrasts were incorporated to identify conserved regulatory modules alongside stage-restricted interactions, effectively capturing the dynamic framework of posttranscriptional regulation throughout developmental transitions.
Statistical analysis
All qPCR experiments were conducted with a minimum of three biological replicates, each analyzed in technical triplicates. The quantification cycle (Cq) values were recorded, and relative gene expression levels were computed using the 2 − ΔΔCq method, with data normalized to the reference genes Actin and U16 small nuclear RNA (snRNA) that were evaluated across developmental stages for stable Cq behavior and minimal inter-stage variation. The normality of the data was assessed via the Shapiro‒Wilk test [40]. Student’s t-test [41] was applied to data exhibiting a normal distribution for pairwise comparisons, whereas the Mann–Whitney U test was employed for nonparametric data. Correlation coefficient and linear regression analyses were performed to compare Cq values across different methods. All data analyses were carried out in R (v.4.5.1).
Results
The study of B. tabaci developmental dynamics using a temporal omics approach revealed two principal ontogenetic stages in its life cycle, characterized by molecular reconfiguration across life stages. Distinct transcriptional profiles were observed across life stages, with a focus on replication and on enhancing detoxification mechanisms in eggs and adults, respectively. These insights provide a deeper understanding of the molecular mechanisms underlying the pest’s developmental plasticity, informing potential pest management. For extended results, please refer to the additional file (Extended File 1), which details mRNA transcriptomic and proteomic analyses, microRNA expression profiles, and the unique molecular blueprints for each developmental stage.
SEM
Scanning electron microscopy (SEM) revealed distinct ultrastructural features across the developmental stages of B. tabaci. The egg phase was observed as an elongated ellipsoid with a smooth chorion, measuring approximately 95–118 μm in length and 78–200 μm in width (Fig. 1a, b). The first instar, or crawler stage, measured approximately 257 μm in body length and presented a dorsoventrally flattened shape with clearly defined segmentation, reflecting its role in dispersal (Fig. 1c). The second instars showed a broader body and enhanced marginal wax secretions (Fig. 1d). In contrast, the third instar, reaching nearly 318 μm, was marked by pronounced dorsal sculpturing and the development of tracheal folds (Fig. 1e). The fourth instar, commonly known as the red-eye nymph, displayed further differentiation in body segmentation, indicating the beginning of pupal development (Fig. 1f). The puparia were notably larger, ranging from 362 to 583 μm in length and nearly 719 μm in maximum width. They were characterized by an oval shape, fused segmentation, and a dense layer of wax on the cuticular surface (Fig. 1g, h). Notably, the anal setae, which measured approximately 177 μm, extended from the posterior margin of the puparial case and served as a crucial diagnostic feature for species-level identification (Fig. 1i).
Fig. 1.
Scanning electron micrographs of B. tabaci developmental stages. Representative images. a and b Egg. c Instar I. d Instar II. e Instar III. f Instar IV. g and h Puparium. i Terminal anal setae, highlighting the morphological detail of the puparium
Stage-resolved multiomic profiling identifies two major developmental transitions in B. tabaci
Integrated profiling of miRNAs, mRNAs, and proteins across egg, instars I–IV, puparium, and adult revealed a structured developmental program with two dominant transition points. Across pairwise stage transitions, approximately 16–34 miRNAs, 1,400–2,300 transcripts, and 500–750 proteins changed significantly, using thresholds of |log2FC|≥ 1 and adjusted p < 0.05 for RNA datasets, and proteomic FDR < 1% for stage-level protein comparisons. The largest system-wide reconfiguration occurred at the egg-to-instar I transition, and a second major shift occurred at the puparium-to-adult transition, whereas intermediate instars showed comparatively narrower remodeling (Figs. 2, 3, 6, Supplementary Figs. 1 and 2; Supplementary Files 2–5). The dataset comprised 59 conserved miRNAs and more than 170 novel candidate miRNAs, indicating that developmental progression is shaped by both conserved regulatory layers and a large repertoire of lineage-enriched small RNAs (Fig. 2f–j; Supplementary Figs. 3; Supplementary Files 2 and 3). Transcriptome and proteome data independently supported these two molecular reconfigurations. The egg-to-instar I comparison yielded the largest DEG set, with about 2,863 DEGs. The instar I-to-II comparison showed about 882 DEGs, mid-instar transitions showed roughly 700–900 DEGs, and the puparium-to-adult transition also showed a major DEG shift (Figs. 3, 4, and 5; Supplementary Files 4 and 6–9). Proteomics identified 874 stage-regulated proteins (Fig. 6a-d), including a shift of more than 200 proteins at egg-to-instar I (Fig. 6e) and more than 300 proteins at puparium-to-adult (Fig. 6f; Supplementary Fig. 4; Supplementary File 5). Together, these data support a model in which hatching and adult emergence represent the two principal molecular reprogramming points in the life cycle (Table 3).
Fig. 2.
Differential expression of known and novel miRNAs across developmental transitions. Volcano plots of significantly differentially expressed miRNAs. a Known miRNAs across Egg → Instar I and b, Instar IV → Adult. c and d Novel miRNAs across the same transitions. Up- and downregulated miRNAs (|log₂FC|≥ 1, padj < 0.05) are highlighted. e Ward clustering of normalized counts across developmental transitions. Top upregulated miRNAs across development: f Strongest upregulated known miRNAs. g Strongest upregulated novel miRNAs. These candidates represent key stage-specific regulators of developmental progression, as shown. h and i Venn diagram of novel and known miRNAs across instars, respectively
Fig. 3.
Differential expression of mRNAs across transitions. Volcano plots of DEGs (|log₂FC|≥ 1, padj < 0.05). a Egg → Instar I. b Instar IV → Adult. c and d Comparative DEG profiles for Egg → Instar I vs Instar I → II and Instar III → IV vs Instar IV → Adult. e Upregulated mRNAs. f Downregulated mRNAs. g Venn diagram of non-significant genes across instars. h Heatmap of normalized counts across developmental transitions. i Ward clustering of normalized counts across developmental transitions
Fig. 6.
Comparative proteomic landscape across instars. a Venn diagram of the shared proteomic profiles. b UpSet plot of overlapping proteins. c Boxplot of imputed peptide intensities. d Heatmap of differentially expressed peptides. Differential expression of proteins across transitions. e Egg → Instar I. f Puparium → Adult. Functional enrichment of proteomes by GO annotation. g Egg. h Adult
Fig. 4.
Gene ontology enrichment of upregulated mRNAs. a Egg → Instar I. b Instar II → Instar III. c Instar IV → Adult
Fig. 5.
Gene ontology enrichment of downregulated mRNAs. a Egg → Instar I. b Instar II → Instar III. c Instar IV → Adult
Table 3.
The developmental atlas of B. tabaci
| Organ System | Instar I | Instar II | Instar III | Instar IV | pupa | Adult |
|---|---|---|---|---|---|---|
| CNS/Neural circuits | Formation & wiring onset – CNS formation, axon guidance, synapse assembly. e.g. LOCUS6761(CNS formation), LOCUS5363(axonogenesis, synapse assembly), LOCUS6798(synapse organization), LOCUS4576(axon midline guidance). | Circuit expansion & refinement – same neural genes remain high, supporting more complex behaviour and sensory integration. | Late refinement – continued high expression of circuit genes; aligning CNS with emerging adult organs (wings, eyes). | Stabilisation – neural architecture largely laid down; circuits link to nearly complete wings, eyes and muscles. | Re-patterning & maturation – pupal remodelling of circuits to adult topology (flight, mating, dispersal). | Functional adult CNS – high synaptic and behavioural activity. e.g. LOCUS6222 (adult locomotory behaviour, synaptic modulation), LOCUS4576 (circuit maintenance). |
| Muscles (body + flight) | Initial myogenesis – somatic/visceral muscle formation for crawling and feeding. e.g. LOCUS13379, LOCUS9028(actin filament organisation), LOCUS5492(regulation of muscle contraction). | Growth phase– existing muscles enlarge and strengthen; contraction genes remain high. | Onset of flight muscle architecture – strong induction of sarcomere/myofibril genes. e.g. LOCUS6348 (actin cytoskeleton, Z-disc, myofibril). | Peak muscle morphogenesis– thoracic/flight muscles heavily built; contraction and sarcomere genes maximal. | Remodelling to adult fibre pattern – pupal remodelling finalises flight muscle architecture. | Functional flight & locomotion – structural genes remain elevated but below morphogenesis peak (maintenance mode). LOCUS5492, LOCUS6348, LOCUS6222 support sustained contraction and movement. |
| Tracheal/Respiratory | Initial tracheal system organisation – tube sealing and barrier formation. e.g. LOCUS2263(open tracheal system, septate junctions, blood–brain barrier). | Scaling with body size – increased gas exchange demand, but no new organ formation. | Metabolic signalling re-tuning (FoxO, HIF-1, cAMP) – optimises tracheal/gas exchange relative to high growth metabolism. | Further tube growth – preparing for adult-scale tracheae feeding thoracic flight muscles. | Adult tracheal network built – branching and diameter tuned for flight oxygen requirements. | High-capacity respiratory function – electron transport & oxidative metabolism high, matching flight activity. |
| Cuticle/Exoskeleton | First functional nymphal cuticle– strong chitin synthesis & catabolism. e.g. LOCUS13278, LOCUS3691, LOCUS2017, LOCUS10150, LOCUS10006(chitin metabolic processes). | Cuticle thickeningwith growth; additional deposition and mild remodelling for next molt. | Progressive strengthening – supports larger body and wing pad outgrowth. | Pre-metamorphic cuticle – robust 4th-instar “pupal-like” cuticle; strong chitin turnover. | Adult cuticle genesis– new exoskeleton built; pupal exuviae discarded. | Hardening & pigmentation – final sclerotisation, colour patterning; stable structural support for wings and muscles. |
| Wings (imaginal disc-derived) | Priming only – discs specified but tiny; wing-hair gene low. | Growth of wing pads – still minimal specific patterning signal. | Wing hair patterning ON – strong upregulation of LOCUS1360 (imaginal disc-derived wing hair organisation). | Wing hair & surface patterning peak – LOCUS1360r emains high; wing membrane & surface structures finalised. | Wing veins & membrane maturation – cuticle & extracellular matrix genes remodel wings to adult form. | Mature wings – LOCUS1360 drops back; wings structurally fixed and used for flight. |
| Eyes/Visual system | Early eye field & morphogenesis– eye development genes turned on. e.g. LOCUS10997, LOCUS4938(eye development, sensory organ morphogenesis). | Primordia build-up – eyes enlarge and differentiate but remain nymph-type. | Strong compound eye morphogenesis– LOCUS10997rises sharply; ommatidial structure elaborated. | Peak visual organ development – LOCUS10997 highest; compound eye structure largely completed. | Refinement & alignment – final pathway remodelling, photoreceptor structuring. | Functional adult compound eye – expression of LOCUS10997 maintained at an intermediate level to support visual function. |
| Gut/Digestive system | Functional digestive tract present; moderate metabolic demand. | Carbohydrate metabolism up – supports high feeding and growth. | Central carbon metabolism (glycolysis, TCA) up – gut and fat body handle intense energy flux. | Detox and transition – preparing for reduced feeding before pupation. | Gut remodelling – reorganisation of epithelium and microbiota/symbionts to adult diet. | Adult feeding physiology– stable digestive function adapted to phloem feeding and vector role. |
| Fat Body/Systemic Metabolism | Basal storage and detox; supports early growth. | Carbohydrate storage & mobilisation – matching growth. | Strong energy metabolism and signalling – FoxO, HIF-1, cAMP; metabolic plasticity. | Lipid accumulation for metamorphosis– fat body prepares energetic reserves. | Redistribution of reserves to developing adult organs. | Adult energy hub – manages energy for flight, reproduction, immune responses. |
| Reproductive organs (gonads) | Priming – spermatogenesis-associated gene appears (e.g. LOCUS2499). | Largely quiescent; somatic growth dominates. | Early gonadal growth; set-up of reproductive primordia. | Pre-gonadal expansion; tissue templates for adult ovaries/testes in place. | Gonad organogenesis – differentiation of germline and somatic structures (ovarioles, testis tubules, accessory tissues). | Functional reproduction– adult oogenesis/spermatogenesis and accessory gland activity fully active. |
| Immune/Symbiont/Virus | Basal immune surveillance. | Mild tuning with growth and diet. | Immune/metabolic signalling coordination. | Transition stage. | Symbiont composition and localisation shift during metamorphosis. | Strong virus/symbiont interface activity – GO: viral RNA replication, symbiont-mediated immune suppression; critical for vector competence. |
The initial molecular surge at hatching establishes the foundational body plan of the crawler
The egg-to-instar I transition represented the most dynamic early-life interval and was characterized by a coordinated burst of transcriptional, post-transcriptional, and proteomic remodeling. At the transcript level, 937 genes were upregulated and 867 were downregulated, with notable induction of COX6B1 (log2FC = + 3.9), ATP synthase F0 subunits (log2FC = + 3.1, p < 0.001), and cuticle protein CPR10 (log2FC = + 4.5, p < 0.001) (Fig. 3a, c-i; Tables 1 and 2; Supplementary File 4). Functional enrichment analysis highlighted oxidative phosphorylation, ribosome biogenesis, and amino sugar metabolism (Figs. 4 and 5; Supplementary Figs. 4 and 5; Supplementary Files 6–9), while proteomic data showed increased abundance of NADH dehydrogenase subunits, ATP synthase complexes, and actin isoforms (Fig. 6a, h; Supplementary File 5). These coordinated changes indicate that hatching is accompanied by strong activation of energy metabolism, along with the structural programs needed for mobility and host interaction.
Table 1.
This table details the first phases of development initiated during the transition from egg to the first instar
| Organ System | Up-regulated Genes | log2FC | GO Biological Process |
|---|---|---|---|
| Central Nervous System (CNS)/Neurogenesis | BEMITA_LOCUS5363 | 4.48 | Axonogenesis (GO:0007409); Synapse assembly (GO:0051965) |
| BEMITA_LOCUS2263 | 3.79 | Glial blood–brain barrier formation (GO:0060857); Septate junction assembly | |
| BEMITA_LOCUS6761 | 3.6 | CNS formation (GO:0021556) | |
| BEMITA_LOCUS6798 | 3.28 | Synapse organization (GO:0050808) | |
| BEMITA_LOCUS4576 | 3.2 | Axon midline guidance (GO:0016199) | |
| Muscle/Myogenesis | BEMITA_LOCUS5492 | 6.44 | Regulation of muscle contraction (GO:0006937) |
| BEMITA_LOCUS13379 | 6.17 | Actin filament organization (GO:0007015) | |
| BEMITA_LOCUS9028 | 4.69 | Actin filament organization | |
| BEMITA_LOCUS12060 | 2.93 | Muscle-related cytoskeletal assembly | |
| BEMITA_LOCUS4576 | 3.2 | Cytoskeletal regulation (muscle) | |
| Tracheal/Respiratory System | BEMITA_LOCUS2263 | 3.79 | Open tracheal system morphogenesis (GO:0035159) |
| Cuticle/Exoskeleton/Chitin | BEMITA_LOCUS13278 | 2.96 | Chitin catabolic process (GO:0006032) |
| BEMITA_LOCUS3691 | 2.9 | Chitin catabolic process | |
| BEMITA_LOCUS2017 | 2.46 | Chitin biosynthetic process (GO:0006031) | |
| BEMITA_LOCUS10150 | 2.15 | Cuticle remodelling | |
| BEMITA_LOCUS10006 | 1.85 | Chitin metabolic process | |
| Sensory Organs (Eyes, Antennae) | BEMITA_LOCUS10997 | 2.98 | Eye development (GO:0001654); Morphogenesis |
| BEMITA_LOCUS4938 | 1.82 | Sensory organ morphogenesis (GO:0009887) | |
| Gonad/Reproductive Axis | BEMITA_LOCUS2499 | 2.53 | Spermatogenesis (GO:0007283) |
| BEMITA_LOCUS4576/LOCUS953 | 1.72 | Transcription regulators linked to reproductive primordia |
Table 2.
This table details the second phase of development, initiated during the transition from the fourth to the fifth instar
| Gene ID | Key GO Biological Process | log2FC | padj | Adult Expression Role |
|---|---|---|---|---|
| BEMITA_LOCUS5492 | Regulation of muscle contraction (GO:0006937) | 5.64 | 0.0021 | Master regulator of muscle contractility. Drives sustained thoracic and abdominal muscle function, including flight beat frequency and host‐settling behaviour. Expression remains high across all instars → Adult retains the full contraction module primed earlier. |
| BEMITA_LOCUS6222 | Adult locomotory behaviour; modulation of chemical synaptic transmission (GO:0008344, GO:0050804) | 2.23 | 0.0053 | Adult locomotion & neural behaviour gene. High expression ensures advanced synaptic firing, walking/climbing agility, mate location, plant-hopping, and dispersal. Strongly linked to behavioural readiness. |
| BEMITA_LOCUS6348 | Flight muscle structure (actin cytoskeleton, Z-disc, myofibril) | 2.51 | 0.012 | Structural pillar of adult flight muscle maintenance. Peaks during Instar 3–4 for morphogenesis, but remains highly expressed in adults to maintain sarcomere alignment, wing-beat stability, and sustained flight endurance. |
| BEMITA_LOCUS10997 | Eye development & photoreceptor morphogenesis (GO:0001654) | 3.98 | 0.0034 | Adult visual system maintenance gene. After strong morphogenesis in Instar 3–4, adults retain high expression for stabilizing phototransduction pathways essential for host detection, leaf selection, mate finding, and navigation. |
This first molecular burst also extended across multiple organ systems. As summarized in Table 1, genes associated with CNS formation and wiring were strongly induced, including BEMITA_LOCUS5363 (axonogenesis and synapse assembly; log2FC = 4.48), BEMITA_LOCUS2263 (glial blood–brain barrier/septate junction assembly; log2FC = 3.79), BEMITA_LOCUS6761 (CNS formation; log2FC = 3.60), and BEMITA_LOCUS6798 (synapse organization; log2FC = 3.28). In parallel, myogenic and cytoskeletal programs were activated through BEMITA_LOCUS5492 (regulation of muscle contraction; log2FC = 6.44), BEMITA_LOCUS13379 (actin filament organization; log2FC = 6.17), and BEMITA_LOCUS9028 (actin organization; log2FC = 4.69), supporting early crawler motility. Additional induction of BEMITA_LOCUS2263 was consistent with tracheal system organization, while BEMITA_LOCUS13278, BEMITA_LOCUS3691, BEMITA_LOCUS2017, and related loci reflected chitin synthesis, catabolism, and cuticle remodeling. Marker genes such as BEMITA_LOCUS10997 and BEMITA_LOCUS4938 further indicated early eye and sensory organ morphogenesis, whereas BEMITA_LOCUS2499 and its associated transcriptional regulators suggested initial priming of the reproductive axis (Table 1). Thus, the hatching transition represents an integrated developmental reset rather than a purely metabolic shift.
At the small-RNA level, this interval was accompanied by pronounced miRNA turnover. The Btab-miR-3791 family was strongly upregulated, with multiple isoforms among the top induced known miRNAs, including Btab-miR-3791-2-3p (log2FC = 9.04), Btab-miR-3791-4-3p (log2FC = 8.61), and Btab-miR-3791-3-3p (log2FC = 8.49) (Supplementary File 2), whereas Btab-miR-989-5p was strongly downregulated (log2FC = − 10.5) (Fig. 2a, e, f, h; Supplementary Fig. 5; Supplementary Files 2 and 3). In addition, 16 egg-biased miRNAs were downregulated, and 12 novel miRNAs were upregulated, including Btab-novel-miR-074a (Fig. 2c; Supplementary Files 2 and 3). qPCR-supported inverse expression between BEMITA_LOCUS14014 and Btab-miR-3791-3 was consistent with a candidate regulatory relationship during this transition (Fig. 7; Supplementary Files 10, 11, and 15). Together, these results indicate that hatching is accompanied by a broad post-embryonic molecular burst that establishes the crawler-stage body plan.
Fig. 7.
Stem‒loop qPCR results. a Validation of miRNA expression. Bar plots of validated miRNAs comparing NGS and qPCR fold changes across developmental transitions. Conserved and novel miRNAs show consistent directionality, confirming the results of the NGS predictions. b miRNA–target repression. Bar plots illustrating validated miRNA–mRNA pairs, including Btab-miR-3791 → BEMITA_LOCUS14014, Btab-miR-2944b → BEMITA_LOCUS7428, and Btab-miR-2944b-1 → BEMITA_LOCUS13425
Intermediate instars consolidate and refine developing systems
In contrast to the two burst-like transitions at hatching and adult emergence, the instar I-to-IV interval was characterized by narrower, yet biologically coherent remodeling. The instar I-to-II transition involved 524 upregulated and 433 downregulated transcripts (Fig. 3c-i; Supplementary Fig. 2) and was enriched for cuticle development, extracellular matrix organization, and carbohydrate processing (Figs. 4 and 5; Supplementary Figs. 4 and 5; Supplementary Files 4 and 6–9). Representative induced loci included chitin synthase 1 (log2FC = + 4.2, p < 0.001) and cuticle protein CPR17 (log2FC = + 3.6, p < 0.001), while proteomic analysis supported this trend through increased cuticular collagens and chitin deacetylases (Supplementary Figs. 2–4; Supplementary File 5). At the post-transcriptional level, 11 novel miRNAs were upregulated, including Btab-novel-miR-054a and Btab-novel-miR-225a (Fig. 1e-i; Supplementary Fig. 1; Supplementary Files 2 and 3), and Btab-miR-2944b showed strong qPCR support (Fig. 7a; Supplementary Fig. 11; Supplementary File 15). These data indicate progressive structural reinforcement during the second instar.
The instar II-to-III transition showed 678 upregulated and 492 downregulated transcripts and was characterized by more pronounced morphogenetic and respiratory remodeling. Upregulated genes included chitinase-like proteins (log2FC = + 3.8, p < 0.001), collagen alpha-1 (log2FC = + 3.4, p < 0.001), and ecdysone-induced E75 (log2FC = + 2.7, p < 0.001), while proteomic analysis identified increased tubulins, cuticular collagens, and respiratory enzymes (Figs. 3c–i, 4, 5 and 6; Supplementary Fig. 2; Supplementary Files 4–9). miRNA remodeling remained evident, with 15 novel miRNAs upregulated, including Btab-novel-miR-027b (log2FC = + 3.4) and Btab-novel-miR-330a (log2FC = + 2.9), which were associated with predicted target sets linked to tracheal and morphogenetic pathways (Fig. 2e-i; Supplementary Figs. 1 and 3; Supplementary Files 2 and 3). Known miRNAs such as Btab-miR-276-3p were also retained within this stage-specific network (Fig. 2e-i; Supplementary Fig. 1 and Supplementary Files 2 and 3), and qPCR-supported inverse expression between BEMITA_LOCUS4129 and Btab-miR-3791-2 was consistent with a candidate regulatory association (Fig. 7a; Supplementary Files 10, 11, and 15). These findings indicate continued organogenesis and respiratory-system refinement during mid-nymphal development.
The instar III-to-IV transition was numerically smaller, with 287 transcripts upregulated and 249 downregulated, but remained biologically distinct (Fig. 3h-i; Supplementary Fig. 2; Supplementary File 4). Enriched categories included cuticle stabilization, carbohydrate derivative metabolism, and oxidative stress response (Figs. 4 and 5; Supplementary Figs. 4–6; Supplementary Files 6–9). miRNA remodeling was again prominent, with strong induction of Btab-novel-miR-018a (log2FC = + 3.6, p < 0.001) and Btab-novel-miR-110a (log2FC = + 2.9, p < 0.001), both associated with predicted structural and extracellular matrix-related target sets (Fig. 2e-i; Supplementary Files 2 and 3). In parallel, Btab-miR-2944b isoforms were downregulated and linked to target categories related to amino acid metabolism and vesicular transport (Fig. 2d, l–m, o; Supplementary Files 2 and 3). Network analysis placed several instar IV-associated miRNAs among the higher-connectivity nodes of the inferred regulatory network (Fig. 8a, b), indicating selective regulatory refinement before pupariation. Overall, the intermediate instars appear to primarily consolidate and refine the developmental systems activated earlier, rather than establishing entirely new molecular states.
Fig. 8.
Integrated multiomics networks of miRNA–mRNA–proteome interactions. a Networks of conserved miRNAs with target mRNAs and proteins. b Networks of novel miRNAs with target mRNAs and proteins. Hubs illustrate stage-specific regulatory hierarchies
This interpretation is further reinforced by the developmental atlas summarized in Table 3, which depicts the progressive, stage-wise maturation of neural circuits, muscles, tracheal organization, cuticle architecture, wing patterning, visual systems, digestive and metabolic capacity, and reproductive tissues across the nymphal instars. For example, neural genes associated with CNS wiring and synapse assembly remain elevated from instar I onward; muscle-related genes transition from crawler-associated myogenesis to flight muscle architecture; and eye-associated genes intensify toward the late nymphal stages as the adult visual system takes shape (Table 3). Therefore, intermediate development is best regarded as a consolidation phase during which previously initiated modules are strengthened, expanded, and integrated.
The second molecular surge at adult emergence facilitates adult physiology and ecological specialization
The puparia and adult emergence intervals defined the second major molecular burst in the B. tabaci life cycle. The puparial stage itself exhibited broad reorganization, with 815 transcripts upregulated and 689 downregulated, and was enriched for proteases, chaperones, and oxidative stress regulators (Figs. 3b, c-i, 4 and 5; Supplementary Files 4 and 6–9). Proteomic analysis independently supported this shift through increased levels of heat shock proteins, proteases, and redox-associated proteins (Fig. 6f, h; Supplementary File 5). At the miRNA level, Btab-novel-miR-199a (log2FC = + 3.5, p < 0.001) and Btab-novel-miR-340a (log2FC = + 3.2, p < 0.001) were among the most strongly induced candidates and were associated with predicted autophagy- and MAPK-related target sets (Fig. b, d, e-i; Supplementary Files 2, 3, 10, and 11). qPCR further supported induction of Btab-novel-miR-199a and Btab-novel-miR-340a, while inverse expression between Btab-miR-2944b-1 and BEMITA_LOCUS13425 was consistent with a candidate remodeling-associated regulatory interaction (Figs. 7 and 8; Supplementary Files 10, 11, 14, and 15).
The puparium-to-adult transition involved 1,147 upregulated and 938 downregulated transcripts and was dominated by adult physiological specialization (Fig. 3b, c–h; Supplementary File 4). Upregulated transcripts included CYP6CX1 (log2FC = + 5.1, p < 0.001), GSTe1 (log2FC = + 4.7, p < 0.001), and OBP4 (log2FC = + 3.9, p < 0.001), which were associated with enriched xenobiotic metabolism, glutathione conjugation, and olfactory transduction pathways (Figs. 4 and 5; Supplementary Files 6–9). Proteomic analysis confirmed the increased abundance of CYPs, GSTs, and odorant-binding proteins in adults (Fig. 6f and h; Supplementary File 5). Adult-biased miRNAs, including Btab-novel-miR-107a (log2FC = + 4.2, p < 0.001) and Btab-novel-miR-307a (log2FC = + 3.9, p < 0.001), occupied high-connectivity positions within inferred networks enriched for predicted targets related to detoxification and chemosensory function (Figs. 2d, e-i, 8a; Supplementary Files 2, 3, 10, and 11). PCA trajectories further showed that adults formed a distinct terminal cluster, farthest from eggs and nymphal instars (Supplementary Fig. 7), underscoring the molecular separation of the adult stage from earlier developmental stages.
Tables 2 and 3 further refine this late developmental burst by showing that adult emergence is accompanied not only by detoxification and sensory activation, but also by functional maturation of locomotion, vision, metabolism, and reproduction. Marker loci retained or elevated in adults included BEMITA_LOCUS5492 (muscle contraction; log2FC = 5.64), BEMITA_LOCUS6222 (log2FC = 2.23), BEMITA_LOCUS6348 (flight-muscle structure; log2FC = 2.51), and BEMITA_LOCUS10997 (eye development and photoreceptor morphogenesis; log2FC = 3.98), indicating that the late burst converts previously assembled developmental modules into an operational adult physiological state (Table 2). In parallel, Table 3 supports maturation of digestive physiology, fat-body metabolic function, gonad activity, and virus/symbiont-associated host interfaces, all of which are consistent with adult dispersal, feeding, reproduction, and vector competence. Together, these data indicate that adult emergence represents a coordinated second molecular burst that enables ecological specialization rather than a simple continuation of nymphal growth.
Conserved and stage-enriched novel miRNAs occupy distinct regulatory positions
Network analysis separated the regulatory architecture into conserved and novel components. In addition, a few top-selected miRNAs were further validated across each instar using stem-loop qPCR (Supplementary Fig. 8), and the overall target predictions are presented in Supplementary Figs. 9 and 10. Conserved miRNAs, including Btab-miR-34 and Btab-miR-2944b, were associated primarily with replication, morphogenesis, and cuticle-associated programs, whereas novel miRNAs dominated later stage-associated modules involved in remodeling and specialization (Fig. 8a, b; Supplementary Files 10 and 11). The stage-specific network structure, together with expression dynamics across transitions (Supplementary Figs. 11–13; Supplementary Files 2–4), supports a division of labor between conserved checkpoint regulators and lineage-enriched modulators, although direct functional validation is available for only a subset of candidate interactions.
Spatial FISH supports stage-restricted co-localization of bta-novel-miR-018a and vitellogenin
To test one predicted regulatory interaction experimentally, whole-mount FISH was performed for Btab-novel-miR-018a and vitellogenin mRNA in late instar IV and adults. Transcriptomic analysis had shown that bta-novel-miR-018a was significantly induced in instar IV (log2FC = + 3.6, p < 0.001) (Fig. 2; Supplementary Fig. 1; Supplementary Files 2 and 3). In instar IV, the FISH study revealed spatial co-localization between miRNA and vitellogenin signals (Fig. 9a–f). In adults, the vitellogenin signal remained detectable; however, the overlapping miRNA signal was no longer observed (Fig. 9e). These data are consistent with a stage-restricted inhibitory relationship operating before adult emergence and relaxed in adults. Because a quantitative co-localization coefficient or fluorescence intensity analysis is not reported, this result is most appropriately interpreted as spatial support for a regulatory interaction rather than definitive proof of direct repression at cellular resolution.
Fig. 9.
Validation of the computationally predicted miRNA (bta-novel-mir-018a)-mRNA (vitellogenin) pair via FISH. a Instar IV mRNA (FAM). b Instar IV miRNA (CY3). c Co-localization of miRNA + mRNA. d, e Localization of vitellogenin on ovarian tissues of instar IV and adult, respectively. f Co-localization of miRNA + mRNA in ovarian tissue, confirming their repressive activity
Egg stalk proteomics defines a morphologically distinct, transporter-enriched reproductive structure
Given earlier physiological evidence that the whitefly pedicel mediates water and solute uptake, we asked whether the B. tabaci egg stalk possesses a distinct transporter-enriched molecular composition. SEM analysis identified a structured egg stalk extending from the posterior pole of the egg (Fig. 10a, b) and inactively inserted into leaf tissue (Fig. 10a, b). Proteomic comparison among eggs with stalks, eggs without stalks, and isolated stalk tissue identified 71 proteins exclusive to the stalk, indicating that the stalk possesses a molecular composition distinct from the egg body (Fig. 11a; Supplementary File 5).
Fig. 10.
SEM characterization of the Bemisia egg stalk. a Representative scanning electron microscopy images showing the stalk attached to the posterior pole of Bemisia eggs. Red-circled regions highlight the proximal and distal structures. The stalk exhibits a tapering architecture (17–51 μm proximally; 9–26 μm distally) with a porous, fibrous distal matrix. The lower-right inset shows a detached stalk fragment with preserved microarchitecture. b Integration of SEM evidence with a cross-sectional leaf illustration showing penetration of the stalk through the cuticle, upper epidermis, and mesophyll. Transporter proteins identified by proteomics—Na⁺/K⁺-ATPase, sugar transporter, phosphate transporter, potassium channel domain–containing proteins, autotransporter domains, and DE-cadherin—are mapped onto the inner stalk region. Arrows indicate predicted nutrient flow from host parenchyma into the developing egg. c-f Developmental expression imaging showing NaPI-III localization in pupae (paired ovaries), adults (ovary region), and co-localization with LOC109039444 in adult tissues (merged channels)
Fig. 11.
Proteomic identification of stalk-specific transporters. a UpSet intersection plot showing the number of unique and shared proteins among egg-with-stalk, egg-without-stalk, and dissected stalk samples. The stalk exhibits a large exclusive cluster (71 proteins), indicating a unique protein repertoire. b Quantitative abundance heatmap of transporter-related proteins across eight biological samples. Sample 8 shows the strongest enrichment of multiple transporter families, consistent with high stalk-specific expression. c Transporter presence/absence matrix across three conditions. Na⁺/K⁺-transporting ATPase α-subunit (highlighted) is uniquely detected in stalk tissue. Additional transporter classes (sugar transporters, phosphate transporters, potassium channel proteins, and ABC transporters) exhibit condition-specific distribution. d Two-dimensional electrophoresis (2-DE) profile of Egg-with-Stalk extract. Protein spots 1–6 (circled) represent distinct, reproducible proteins associated with predicted transporter molecular weight ranges. e, Cloning and expression of stalk-associated transporters. f Microbial synthesis of dsNaPi-III and dsGFP. g Number of eggs laid in treatment and control females. h Percent of egg hatched. i Expression results of NaPi-III treatments and controls. j and k Egg laid on control. l and m Eggs laid on treatment
The stalk proteome was enriched in transport-associated proteins, including Na +/K + pumps, sugar transporters, phosphate transporters, potassium channels, ABC transporters, and EGF-like domain proteins (Figs. 10a and 11a, b). Fluorescent imaging further demonstrated localization of NaPi-III to the egg stalk (Fig. 10c-g), with additional expression observed in puparial ovaries and adult abdominal tissues, as well as co-localization with LOC109039444 within adult ovarian tissues (Fig. 10f). These results indicate that the stalk possesses a transporter-enriched molecular profile distinct from the egg body, which contributes to nutrient absorption in eggs in both early and post-embryonic development.
Two-dimensional gel analysis further supported the presence of transporter-associated stalk protein signatures, with distinct spots observed in stalk-containing material (Fig. 11a-d). MALTI/TOF–MS results for these distinct spots further support the interpretation that the stalk is enriched with transporter-rich appendage as identified by LC–MS/MS. However, these data suggest that the transport-associated molecular specialization is not direct evidence of nutrient uptake kinetics, as direct flux measurements of the egg stalk were not performed in this study.
Two transporter candidates, NaPi-III and LOC109039444, were selected for validation. Both transcripts were cloned successfully (Fig. 11a–c), and heterologous expression of NaPi-III in E. coli produced a distinct band approximately 55 kDa in size, aligning with the predicted protein dimensions in LC–MS/MS and MALTI-TOF/MS (Fig. 11d). Functional perturbation assays were performed across three independent biological replicates using untreated control, dsGFP negative control, and dsNaPi-III treatments (Fig. 11e–m). In the control and dsGFP groups, a total of 23 and 30 eggs were laid (Fig. 11g), of which 18 and 20 hatched, corresponding to mean hatch rates of 78.3 ± 6.2% and 66.7 ± 3.2%, respectively (Fig. 11h). In contrast, in the dsNaPi-III group, 24 eggs were laid but only 3 hatched (Fig. 11g), giving a mean hatch rate of 12.5 ± 1.0% (Fig. 11h). Thus, oviposition was only modestly reduced in the dsNaPi-III treatment relative to the untreated control, whereas hatching declined sharply compared with both the untreated and dsGFP controls. Consistent with this phenotype, eggs derived from dsNaPi-III-treated adults frequently showed collapsed chorions, flattened shape, and other marked developmental abnormalities (Fig. 11j, k), whereas eggs from the control groups retained the typical healthy ellipsoidal morphology (Fig. 11l, m). In parallel, NaPi-III RT-qPCR showed that the target-gene Cq values were higher in the dsNaPi-III group (27.01 ± 0.13) than in the untreated control (25.10 ± 0.10) and dsGFP control (25.28 ± 0.18), consistent with reduced NaPi-III transcript abundance after dsRNA treatment (Fig. 11i). Together, these data support a functional association between NaPi-III and normal egg viability within the stalk-associated transporter system.
Integrated evidence identifies two major molecular transition points in B. tabaci development
Taken together, the data identify hatching and adult emergence as the two major molecular transition points in B. tabaci development. The first major transition, at hatching, is characterized by broad metabolic and structural activation, including 937 induced and 867 repressed transcripts (Tables 1, 2 and 3, Fig. 3; Supplementary File 4), strong miRNA turnover (Fig. 2; Supplementary Files 2 and 3), and a major proteomic shift (Fig. 6; Supplementary File 5). Intermediate instars show narrower but still stage-specific remodeling (Figs. 3, 4, 5, 6, 7 and 8; Supplementary Files 2–15), while the second major transition at adult emergence includes strong transcriptomic and proteomic reorganization associated with adult physiology (Fig. 2, 3, 4, 5 and 6; Supplementary Files 2–9). In parallel, the egg stalk emerges as a morphologically distinct and transporter-enriched structure containing 71 exclusive proteins (Figs. 10 and 11; Supplementary File 5) and is functionally linked to egg viability through NaPi-III-associated phenotypes (Fig. 11). Overall, these results provide quantitative support for two principal developmental reprogramming phases and identify the egg stalk as a specialized reproductive structure with strong molecular evidence for transport-related function.
Candidate reference miRNAs for developmental-stage normalization
To establish reliable references for studying B. tabaci, we carefully examined both novel and known miRNAs across all life stages (egg, nymphal instars I–IV, puparium, and adult). Using NGS and stem‒loop qPCR, we found that six miRNAs presented stable expression levels with minimal fluctuations throughout these stages. These include three novel miRNAs, Btab-novel-miR-316a, Btab -novel-miR-376a, and Btab -novel-miR-254a, as well as three known miRNAs: Btab-miR-276-3p, Btab-bantam-1, and Btab-Mir-1 (Fig. 7a, Supplementary Fig. 15, and Supplementary File 15). All six candidates showed minor variation in Cq and log2FC values, indicating no stage-specific increases or decreases in expression. Notably, Btab-miR-276-3p and Btab-bantam-1 are conserved across various insect species and have been identified as reliable regulators in other studies. This finding supports their use as universal reference controls. By adding novel miRNAs, we can expand normalization options, which is especially useful because conserved miRNAs can sometimes show minor variations under certain conditions. These six miRNAs serve as useful references for gene expression studies in B. tabaci. The consistent levels of these miRNAs across all instars provide a solid foundation for accurately normalizing miRNA profiles in stage-specific experiments.
Discussion
An integrative multi-omics analysis of B. tabaci revealed that the most pronounced developmental reorganization centers around hatching and adult emergence. Rather than defining a new developmental paradigm, these data position B. tabaci within a broader insect pattern in which early post-embryonic activation and the juvenile-to-adult transition are accompanied by extensive transcriptomic and proteomic remodeling. The principal advance of this study is the stage-resolved integration of miRNAs, mRNAs, and proteins across the whitefly life cycle, thereby providing a valuable developmental resource for this important pest species. Consistent with this pattern, transcriptome-wide analyses in Tribolium castaneum have identified major shifts between late eggs and early larvae, and again between late pupae and early adults, with DNA replication, cell-cycle, and endocrine pathways among the most strongly remodeled categories [12]. Furthermore, the broader evolutionary conservation of the juvenile-to-adult transition is supported by studies identifying E93 as a conserved adult-specifying factor in both hemimetabolous and holometabolous insects [13]. Comparable adult-biased transcriptional specialization has also been documented in other insects undergoing metamorphosis, including butterfly head transcriptomes in which more than half of expressed genes differed between larval and newly eclosed adult stages, with strong enrichment of sensory-function genes in adults [42].
At the cellular level, conserved miRNAs impose temporal checkpoints, whereas novel miRNAs refine physiological and structural programs, leading to transcriptional changes that are ultimately reflected in proteomic profiles required for the expression of specific phenotypes. We observed that conserved miRNAs, particularly miR-34 and miR-252b, were abundant in eggs but diminished in instar I. These decreases coincided with the upregulation of transcripts involved in replication licensing, ribosome biogenesis, and chromatin remodeling, supporting a model in which maternal miRNAs suppress the premature translation of growth-promoting factors until after hatching. Following this, the release of repression initiates the cell cycle and biosynthesis. This function aligns with previous research in the silkworm Bombyx mori, where miR-34 has been shown to regulate larval growth and wing morphogenesis by repressing cuticle protein genes in coordination with ecdysone signaling [43]. Furthermore, recent findings suggest that miR-34-5p mediates 20-hydroxyecdysone-induced autophagy in the fat body of B. mori by targeting Atg1, linking hormonal signaling to cellular remodeling [44]. These findings collectively affirm the role of conserved miRNAs as developmental checkpoints that connect endocrine signals with cellular proliferation.
The current study on mRNA and proteome dynamics revealed high concordance across developmental stages. For example, genes related to oxidative phosphorylation and structural proteins peak during instar I, whereas cuticular and collagenous networks dominate instars II and III. During pupation, proteolytic and chaperonin activities increase, with detoxification enzymes and odorant-binding proteins becoming prominent in adults. This close alignment suggests that transcriptional programs are largely realized at the protein level, with miRNAs providing fine-tuning in both amplitude and timing. This pattern is consistent with developmental proteome analyses in D. melanogaster, which documented extensive proteome remodeling across the life cycle and showed that stage-specific molecular programs are reflected at the protein level [45]. Similar observations in B. mori [46] emphasize that stage-specific alignments exist for metabolic and structural pathways. Our findings reinforce the idea that core developmental modules are reliably translated into proteomic outputs, whereas any discrepancies likely arise from rapid posttranslational regulation during remodeling or from tissue-specific translation responses. Another significant finding is the distinct roles played by conserved versus novel miRNAs throughout development. Conserved miRNAs, such as Btab-miR-34 and Btab-miR-2944b, are instrumental in guiding early transitions, whereas novel candidates (bta-novel-miR-307a, 352a, and 107a) have emerged as crucial regulators during adult stages. This paradigm reflects evolutionary models that propose a foundational core of conserved insect miRNAs with extensive developmental roles, whereas novel miRNAs are often adapted for specific lineage requirements. In B. tabaci, adult-specific novel miRNAs target detoxification enzymes and pathways involved in sensory receptors, which are vital for defense against xenobiotics and host plant recognition. Similar developmental patterns in cockroaches demonstrate how novel miRNAs regulate cuticular and morphogenetic transcripts, highlighting the need to balance maintaining established developmental processes with adapting to ecological demands [47]. A key limitation is that most proposed miRNA–mRNA interactions are inferred from stage-specific expression, computational prediction, inverse associations, and co-localization, not direct biochemical validation. Thus, the highlighted miRNAs are candidates and hypotheses within the developmental network. Confirming these requires dual-luciferase assays, mutant analyses, and in vivo experiments to verify binding and causal roles.
During the transition from the instar stage to the pupal stage, we observed an enrichment of proteomic factors related to autophagy, protease activity, and chaperones, which align with novel miRNAs predicted to target MAPK and oxidative stress-related transcripts. This observation underscores a conserved axis in which hormone signaling, miRNA activity, and autophagic processes coordinately regulate tissue remodeling. Such integrative mechanisms have been well characterized in B. mori, where miR-34 directly connects 20E signaling to the autophagy machinery [44]. This synergistic interaction likely facilitates the synchronized degradation of larval tissues while simultaneously constructing adult structures, reflecting a deeply rooted strategy across holometabolous insect species. Notably, Adult-biased transcriptional specialization is seen in other metamorphosing insects; butterfly head transcriptomes show larval–adult differential expression, with adult enrichment of sensory genes [42]. These gene families have been well documented as pivotal in insecticide resistance and host plant discrimination in B. tabaci and closely related pests [48]. Additionally, GSTs are expressed in chemosensory organs and are involved in both odorant clearance and detoxification processes [49, 50]. The positioning of novel miRNAs upstream of these regulatory modules underscores the importance of posttranscriptional mechanisms, which serve as vital tools for rapid ecological adaptation.
Our findings extend prior physiological evidence for pedicel-mediated uptake of nutrients in whiteflies by defining a transporter-enriched molecular landscape of the B. tabaci egg [14]. The selective enrichment of the Na⁺/K⁺-ATPase α-subunit, together with specific sugar, phosphate, and potassium transporters, underscores an active nutrient-uptake mechanism mediated by ion gradients, aligning fully with the recognized functions of these pumps and channels in insect epithelial transport [16]. This molecular evidence extends prior physiological studies, which demonstrated significant water and solute uptake by whitefly eggs from host tissue via the pedicel [51], by pinpointing the transporters likely responsible for this process. Distinct proteomic profiles among stalk, egg-with-stalk, and egg-without-stalk samples, combined with remarkable transporters, confirm that the stalk serves as a functionally specialized tissue. The developmental localization of NaPI-III and LOC109039444, exclusively in the stalk during the egg stage, and their reorganization within ovarian tissues of pupae and adult domains further suggests these transporters contribute to a conserved resource-allocation mechanism. Comparable regulatory dynamics of transporter expression across insect life stages have been documented as adaptive strategies for shifting physiological demands [18]. Since dsRNA was administered via adult feeding, the transfer to the ovaries and egg tissues was inferred rather than directly demonstrated. Future investigations should quantify NaPi-III knockdown within the ovary, egg body, and stalk tissue.
From an evolutionary standpoint, the stalk emerges as a novel innovation—a conduit that provides the embryo with direct access to host plant nutrients via a specialized transporter system. Evolutionary diversification of ion transport genes, such as Na⁺/K⁺-ATPase, is well established among insects facing varied ecological pressures [17], and our data support their adaptive recruitment within this specialized reproductive organ. Notably, the centrality of these transporters in reproduction is reinforced by previous observations that Na⁺/K⁺-ATPase knockdown reduces egg production in insects [19]. In conclusion, this study presents the first molecular and proteomic characterization of egg-stalk specialization in B. tabaci, providing candidate transporter components that may underlie the previously described pedicel-associated uptake function. By elucidating its transporter profile and developmental regulation, we provide a mechanistic framework for understanding embryo–host interactions and highlight a novel target for strategies aimed at whitefly reproduction.
Conclusion
This study provides a stage-resolved multiomics atlas of B. tabaci development, showing that the largest molecular reorganization events occur at hatching and adult emergence. The work, therefore, extends conserved insect developmental patterns to an important hemipteran pest while providing candidate stage-biased regulatory modules for future mechanistic investigation. The initial phase involves conserved miRNAs acting as developmental checkpoints to prevent premature proliferation, whereas the later phase involves novel miRNAs that coordinate metamorphic changes and ecological adaptations. RNA sequencing insights are reflected at the proteome level, confirming that developmental regulation is achieved through a precise hierarchical framework. In addition to enhancing our understanding of whitefly biology, these findings underscore a broader evolutionary principle: conserved miRNAs maintain developmental integrity, whereas novel regulatory elements are often repurposed for lineage-specific adaptations, such as detoxification and chemosensation. This dual framework offers both stability and adaptability, explaining how B. tabaci attains ecological versatility and environmental resistance as a pest. From a practical standpoint, identifying stage-biased miRNA hubs and their downstream targets offers promising opportunities for innovative RNA-based control strategies to selectively target B. tabaci development. This research reveals the intricate hierarchy of insect developmental biology, illustrating how multilayered molecular regulation synchronizes growth, morphogenesis, and ecological competence. This integrative molecular atlas not only advances our knowledge of insect evolution but also lays the groundwork for targeted interventions against significant agricultural pests.
Supplementary Information
Acknowledgements
The authors sincerely thank the Department of Agricultural Entomology and the Department of Plant Molecular Biology and Bioinformatics, Tamil Nadu Agricultural University, Coimbatore, Tamil Nadu, India, for providing research facilities. P.K. acknowledges 'National Fellowship for Other Backward Classes (NFOBC)', by the University Grants Commission -Council of Scientific and Industrial Research (UGC-CSIR), Government of India, for providing the fellowship for his doctoral studies.
Authors’ contributions
M.M. and P.K. conceived the study and supervised the project. P.K., M.M., S.T., S.P.S., B.V., J.M., and S.S. conducted the experiments, collected the samples and analysed them. P.K., Y.K., J.M., and S.S. implemented software and performed analyses. M.M. and J.M. curated data sets. P.K., M.M., S.T., Y.K., S.P.S., B.V., J.M., and S.S wrote and edited the manuscript. All authors discussed the results and implications.
Funding
The research did not receive any funds.
Data availability
The analysis results are presented in the manuscript, while additional detailed results and supporting information are provided in the Supplementary Files 1–15. The raw sequencing reads derived from miRNA and mRNA sequencing of B. tabaci generated in this study have been deposited in the NCBI database under the bio project PRJNA1335214, and the proteomics data have been submitted to the jPost proteome repository under JPST004134 (https://repository.jpostdb.org/entry/JPST004134). The complete cloned gene sequences were submitted to NCBI under the following accessions: PX637796 and PX637797.
Declarations
Ethics approval and consent to participate
Not applicable.
Consent for publication
Not applicable.
Competing interests
The authors declare no competing interests.
Footnotes
Publisher’s Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Contributor Information
Prakash Kolanchi, Email: prakash280899@gmail.com.
Murugan Marimuthu, Email: muruganmarimuthu@tnau.ac.in.
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Supplementary Materials
Data Availability Statement
The analysis results are presented in the manuscript, while additional detailed results and supporting information are provided in the Supplementary Files 1–15. The raw sequencing reads derived from miRNA and mRNA sequencing of B. tabaci generated in this study have been deposited in the NCBI database under the bio project PRJNA1335214, and the proteomics data have been submitted to the jPost proteome repository under JPST004134 (https://repository.jpostdb.org/entry/JPST004134). The complete cloned gene sequences were submitted to NCBI under the following accessions: PX637796 and PX637797.