Evolution of circadian clock and light-input pathway genes in Hemiptera

Evolution of circadian clock and light-input pathway genes in Hemiptera | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Evolution of circadian clock and light-input pathway genes in Hemiptera Vlastimil Smykal , Hisashi Tobita , David Dolezel doi: https://doi.org/10.1101/2025.01.17.633578 Vlastimil Smykal 1 Biology Centre of the Czech Academy of Sciences; České Budějovice , 37005, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: v.smykal{at}entu.cas.cz david.dolezel{at}entu.cas.cz Hisashi Tobita 1 Biology Centre of the Czech Academy of Sciences; České Budějovice , 37005, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site David Dolezel 1 Biology Centre of the Czech Academy of Sciences; České Budějovice , 37005, Czech Republic Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: v.smykal{at}entu.cas.cz david.dolezel{at}entu.cas.cz Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract Circadian clocks are timekeeping mechanisms that help organisms anticipate periodic alterations of day and night. These clocks are widespread, and in the case of animals, they rely on genetically related components. At the molecular level, the animal circadian clock consists of several interconnected transcription-translation feedback loops. Although the clock setup is generally conserved, some important differences exist even among various insect groups. Therefore, we decided to identify in silico all major clock components and closely related genes in Hemiptera. Our analyses indicate several lineage-specific alterations of the clock setup in Hemiptera, derived from gene losses observed in the complete gene set identified in the outgroup, Thysanoptera, which thus presents the insect lineage with a complete clock setup. Nilaparvata and Fulgoroidea, in general, lost the (6-4) photolyase, while all Hemiptera lost FBXL3, and several lineage-specific losses of dCRY and jetlag were identified. Importantly, we identified non-canonical splicing variants of period and m-cry genes, which might provide another regulatory mechanism for clock functioning. Lastly, we performed a detailed reconstruction of Hemiptera’s light input pathway genetic repertoire and explored the horizontal gene transfer of cryptochrome-DASH from plant to Bemisia . Altogether, this inventory reveals important trends in clock gene evolution and provides a reference for clock research in Hemiptera, including several lineages of important pest species. ○ Evolution of clock genes (including light input pathway) was reconstructed in Hemiptera ○ New m-cry and per splicing variants were identified in certain species ○ A unique horizontal gene transfer of plant/fungal CRY-DASH was found in Bemisia ○ Clock setup was identified for pests: Nilaparvata , Bemisia , Halyomorpha , and aphids ○ Future clock research directions in Hemiptera are proposed 1. Introduction Life on Earth is heavily affected by periodic changes in day and night. Consequently, nearly all organisms evolved circadian clock, an internal oscillatory mechanism measuring 24 hours (from Latin circa… approximately, diem… day), which helps them to anticipate daily changes. At a molecular level, the circadian clock of animals in general and bilateral animals in particular (i.e. mammals and insects) rely on negative transcription/translation feedback loops (TTFL) of homologous components. At least 20 clock components may be considered circadian clock proteins participating in TTFLs of animal clocks. These proteins include transcription factors, several kinases, phosphatases, proteins responsible for specific degradation pathways, etc. Although the majority of clock components are conserved across animals, certain important differences can be found. Thanks to genetic tools and decades of effort by hundreds of scientists, the fruit fly Drosophila melanogaster has become the best model for the invertebrate circadian clock ( Hall 2003 ; Hardin 2011 ; Ozkaya and Rosato 2012 ; Dolezel 2023 ). CYCLE (CYC) and CLOCK (CLK), two proteins from the basic helix-loop-helix (bHLH) Per-ARNT-Sim (PAS) family ( Tumova et al., 2024 ), serve as positive elements of the feedback loop and drive mRNA expression of genes containing E-box elements in the promoter, such as period ( per ) and Drosophila -type timeless ( d-tim ; for the mammalian-type tim , m-tim , see text on the light input below) ( Allada et al., 1998 ; Rutila et al., 1998 ). PER and dTIM proteins accumulate and enter the cell nucleus, resulting in inhibition of the transcriptional activity of the CYC-CLK dimer ( Glossop et al., 1999 ) ( Fig. 1 ). Although PER protein can form a homodimer ( Landskron et al., 2009 ), the major PER-stabilizing partner is dTIM ( Sehgal et al., 1994 ). Interaction between both proteins is key for their subcellular localization, including nuclear localization signals (NLS) and nuclear export signals (NES) (Saez et al., 1996; Singh et al., 2019 ; Giesecke et al., 2023 ). Both PER and dTIM are phosphorylated/dephosphorylated by several kinases and phosphatases contributing to their subcellular localization and stability, ultimately leading to their ubiquitination and degradation by the proteasome ( Grima et al., 2002 ; Ko et al., 2002 ). Experiments on Drosophila PER functionally grouped phosphorylation sites into several phosphocluster regions, which based on their phosphorylation level determine PER stability and inhibitory potency and modulate temperature compensation phenotype, forming thus a phospho-switch to balance diverse PER functions throughout the day-night cycle ( Kivimäe et al., 2008 ; Joshi et al., 2022 ). One of the phosphoclusters, the N-terminal phosphodegron, is responsible for PER ubiquitination and degradation in the proteasome ( Chiu et al., 2008 ; Kivimäe et al., 2008 ). It includes Serine 44,45 and 47 ( Chiu et al., 2008 ; Joshi et al., 2022 ), which of S47 is phosphorylated by DOUBLETIME (DBT) kinase and together with nearby phosphorylated sites, phospho-S47 generates a high-affinity atypical binding site for SUPERNUMERARY LIMBS (SLIMB) ( Chiu et al., 2008 ), an F-box protein targeting PER for proteasomal degradation ( Grima et al., 2002 ; Ko et al., 2002 ). Download figure Open in new tab Fig. 1. (A) Simplified summary of the most variable components in insect clockwork models. The generally conserved positive components, CLOCK (CLK) and BMAL/CYCLE (CYC) drive the expression of negative feedback loop components. Adapted from Doležel 2023. (B) In Drosophila , key negative components include PER and dTIM. The light-induced degradation of these components requires dCRY, JET, and BRWD3. (C) In addition to Drosophila components, the monarch butterfly possesses mammalian-type CRY (mCRY), which acts as the primary repressor. (D) The honeybee has lost dTIM, dCRY, and JET. (E) The clock in the linden bug relies predominantly on mCRY and PER, with some contribution from dTIM. As the most thoroughly studied clock genes, per and d-tim , can illustrate the fine-tuning role of alternative splicing in circadian clock regulation. At low temperatures, efficient splicing of an intron located in the 3’ untranslated region (UTR) of per results in advance accumulation of PER protein, which further manifests in advanced evening activity typical for fruit flies at low temperatures ( Majercak et al., 1999 , 2004 ). A complex splicing pattern has been reported for d-tim alternative splicing. For example, alternative retention of an intron located in the center of the gene at high temperatures results in mRNA that is likely destroyed by a non-sense mediated decay mechanism (NMD) and can only encode unstable and nonfunctional protein ( Martin Anduaga et al., 2019 ; Foley et al., 2019 ; Shakhmantsir et al., 2018 ). This splicing might be one of many adjustments cumulatively resulting in a temperature-compensated clock in Drosophila . The circadian oscillation is further regulated by the action of Clock-Interacting Protein Circadian (CIPC) and CLOCKWORK ORANGE (CWO). First, when CLK-CYC is released from E-boxes, CWO binds to E-boxes and reinforces the repression ( Zhou et al., 2016 ). At the same time, CWO simultaneously represses the expression of Cipc , which itself is a repressor of CLK-CYC transcription ( Rivas et al., 2021 ). Thus, while CWO represses CLK-CYC, it also activates their transcription later by repressing their repressor, CIPC, altogether explaining the seemingly contradicting roles of simultaneous activator and repressor ( Kadener et al., 2007 ; Richier et al., 2008 ). Another group of TTFL involves basic leucine zipper (bZIP) proteins VRILLE (VRI), PAR DOMAIN PROTEIN 1 (PDP1), and KAYAK (KAY). Their genes, specifically certain isoforms, contain E-boxes that are recognized by the CLK-CYC dimer and drive the periodic mRNA transcription. First, the mRNA of vrille ( vri ) peaks, whereas the Pdp1-epsilon isoform reaches its maximum several hours later, and a similar delay is observed in protein abundance. VRI serves as a transcription repressor that recognizes D-boxes (also known as V/P boxes), cis-regulatory sequences located in the Clk promoter ( Blau and Young, 1999 ; Cyran et al., 2003 ). The role of VRI is gradually mitigated by two additional bZIP proteins. KAY, a homolog of mammalian FOS, directly binds VRI and inhibits its repressive role ( Ling et al., 2012 ); whereas PDP1 recognizes and binds to the same DNA sequence. However, PDP1 serves as a transcriptional activator; thus, the actions of the three bZIP proteins described here result in periodic expression of Clk mRNA that is in antiphase to per and d-tim oscillation in Drosophila ( Cyran et al., 2003 ). The mammalian clock relies on a TTFL with RORα and REV-ERBα, two orphan nuclear receptors cyclically expressed by CLK-BMAL (Brain and Muscle ARNT-like 1, a mammalian homolog of CYC). Similarly, Drosophila clock machinery involves nuclear hormone receptors E75 (homolog of REV-ERBα) and UNFULFILLED (UNF) ( Jaumouille et al., 2015 ). Interestingly, the role of UNF seems to be a Drosophila -specific clock idiosyncrasy, as this protein has not been identified as a clock component in other insect models. Instead, HR3 (homolog of RORα) was identified by RNA interference in an early-branching insect species, the firebrat Thermobia domestica ( Kamae et al., 2014 ). TAIMAN (TAI), a bHLH-PAS protein also known as steroid receptor coactivator (SRC) or Nuclear Receptor Coactivator (NCoA), was recognized as a new clock component in two distant insect lineages: true bugs and cockroaches ( Smykal et al., 2023 ). TAI, initially identified as a developmental gene in a forward genetic screen ( Bai et al., 2000 ), participates in numerous developmental processes, including metamorphosis, reproduction, and adult diapause ( Smykal et al., 2014a ; Smykal et al., 2014b ; Hejnikova et al., 2022 ). Its roles involve participation in juvenile hormone and ecdysteroid receptions, but its function in the circadian clock seems to be independent of either of these hormones. Importantly, the mammalian ortholog SRC2 participates in mouse circadian rhythmicity, while SRC3 is involved in ultradian metabolic rhythms, which further underscores the evolutionarily conserved participation of TAI in the clock ( Stashi et al., 2014 ; Meng et al., 2022 ). The role of TAI in the Drosophila circadian clock remains elusive. As already illustrated regarding nuclear receptors ( Kamae et al., 2014 ; Jaumouille et al., 2015 ), there are specific modifications in the clock setup related to particular insect lineages. Additional major lineage-specific changes involve the dTIM protein and its partners. Similar to the mammalian clock, dTIM has been lost in Hymenoptera ( Rubin et al., 2006 ) and independently in termites ( Kotwica-Rolinska et al., 2022a ). Notably, this gene loss is further accompanied by the loss of Drosophila -type cryptochrome , d-cry (in the Drosophila literature often annotated as cry , in some evolutionary comparisons annotated as cry1 or cryI ), a gene encoding protein related to DNA photolyases that lacks the DNA-repair activity ( Yuan et al., 2007 ; DeOliveira and Crane, 2024 ). Another homolog of DNA photolyases, the mammalian-type cry , m-cry (annotated as cry2 or cryII in some evolutionary comparisons), has been lost in Drosophila . The mCRY protein, first identified as a major component of the clock in vertebrates ( Kume et al., 1999 ; Putker et al., 2021 ), is a key component of clock machinery in numerous insects, including the monarch butterfly, cockroaches, and true bugs (Ikeno 2011a; Bazalova et al., 2016 ; Zhang et al., 2017 ; Werckenthin et al., 2020 ; Kotwica-Rolinska et al., 2022a ). Although dCRY contributes to the clock oscillation and robustness in Lepidoptera and to some extent even in Drosophila ( Dolezelova et al., 2007 ; Iiams et al., 2024 ; Tobita and Kiuchi, 2024 ), particularly in the peripheral oscillator ( Collins et al., 2006 ), its major role in the fruit fly is in the light-mediated entrainment, where dCRY serves as a deep brain photoreceptor ( Emery et al., 2000 ). Upon blue light illumination, dCRY directly interacts with dTIM, and both proteins are targeted for proteasomal degradation when JETLAG (JET) and BRWD3/Ramshackle play key roles ( Ceriani et al., 1999 ; Koh et al., 2006 ; Peschel et al., 2009 ; Ozturk et al., 2013 ). While the latter is a developmental gene ( D’Costa et al., 2006 ), JET has been lost in several insect lineages ( Kotwica-Rolinska et al., 2022a ), and often, but not always, the loss accompanies either the loss of dCRY or major modification of dTIM ( Bullo et al., 2024 ). Interestingly, dCRY-independent light input affecting dTIM has been reported for the QUASIMODO (QSM), a membrane-anchored Zona pellucida domain protein ( Chen et al., 2011 ). The detailed mechanism of QSM action in the Drosophila clock remains elusive and so is the role of this protein in the clock of other species. In addition to dCRY-dTIM entrainment, light information accesses the Drosophila clock via canonical visual opsins ( Saint-Charles et al., 2016 ). These photoreceptors are found in the compound eyes, ocelli, and the Hofbauer-Buchner eyelet, which is located underneath the compound eye and projects to the brain pacemaker (Helfrich-Förster et al., 2001; Helfrich-Förster 2019 ). Their phototransduction cascade includes two phospholipases C-β encoded by the no receptor potential A ( norpA ) gene and PLC-β encoded by the Plc21C gene ( Helfrich-Forster et al., 2001 ; Ogueta et al., 2018 ). In crickets, compound eyes are the only circadian photoreceptors. Both the opsins, particularly the green-sensitive one, and cooperation between cryptochromes and c-fos mediate photic entrainment of the circadian clock ( Komada et al., 2015 ; Kutaragi et al., 2018 ). Similarly, the retinal cells in the compound eyes are key for the light input into the photoperiodic timer in the bean bug Riptortus pedestris ( Xi et al., 2017 ). This contrasts with the localization of the light-sensitive regions to the brain in aphids, as was elegantly shown using a capillary-focused light source ( Lees 1964 ). However, the actual photoreceptor is not clearly pointed because aphids possess dCRY, dTIM, and opsins ( Cortés et al., 2010 ; Collantes-Alegre et al., 2017). Thus, even more closely related groups (such as aphids and bean bugs) may differ in the anatomy of their light input. As briefly illustrated above and in literature ( Tomioka 2014 ; Tomioka and Matsumoto 2015 ; Tomioka and Matsumoto 2019 ; Kotwica-Rolinska et al., 2022a ; Dolezel 2023 ), certain important differences exist in the clock setup among various insect lineages. Hemiptera is a large insect order and together with Psocodea and Thysanoptera form Hemipteroid insects (Paraneoptera) ( Johnson et al., 2018 ). This large assemblage contributes to more than 10% of insects’ diversity and with numerous pests and disease vectors they are important both from basic research and practical perspectives. The circadian clock of Hemiptera was explored to a different extent in some species. In Rhodnius prolixus , the anatomy of the clock was studied in the neuroanatomical context and from the developmental perspective ( Vafopoulou et al., 2007 ; Vafopoulou and Steel, 2012 ). Similarly, the localization of circadian clock products was assessed using in situ hybridization ( m-cry , d-cry , per and d-tim transcripts) and immunohistochemistry (dCRY and PER antibodies) in the pea aphid A. pisum, and analyzed in the context of possible seasonality-relevant outputs, such as insulin-like proteins ( Colizzi et al., 2021 ; Barberà et al., 2017 , 2022 ). Furthermore, splicing isoforms were analyzed with temporal resolution under both short and long photoperiods, however, no association of specific transcript variants to a particular aphid strain or photoperiod could be identified ( Barberà et al., 2017 ). At a functional level, the circadian clock genes were analyzed in two hemipteran model species, the bean bug R. pedestris and the linden bug P. apterus . In both species, systemic RNA-mediated interference (RNAi) is routinely used. Furthermore, in the linden bug, CRISPR-Cas9 gene editing was established and used to fully explore the role of several clock genes ( Kotwica-Rolinska et al., 2019 , 2022a , 2022b ). Given the above-described complexity of circadian clocks in insects, the inventory of clock genes in Hemiptera promises to identify important changes in the clock setup of several agricultural pests including aphids, psyllids, true bugs, and brown plant hoppers. Revealing lineage-specific traits unique to this diverse group of pests, or identifying more general patterns within Hemiptera, is essential, as these traits may influence their behavior, including the daily activity cycles, regulation of seasonality, and reproduction. Additionally, these genes are likely involved in migratory behaviors, which can significantly affect how many pest species move between habitats, especially in response to seasonal variations. Therefore, a comprehensive inventory of circadian clock genes is necessary to elucidate these functional relationships, paving the way for more effective pest management strategies that consider the intricate biological and ecological interplay dictated by circadian mechanisms. 2. Methods and materials 2.1 Dataset and gene discovery We used a similar approach to identify genes in selected insect lineages as in Smykal et al. (2020) . In brief, the initial steps of gene identification involved a BLASTP search in the protein database, a TBLASTN search in transcriptome shotgun assemblies (TSA), and we also used keywords if the genome/protein dataset was annotated. Importantly, several hits from the same species (usually more than five) were retrieved for the initial analysis. All retrieved sequences were first aligned using MAFFT, and fast tree analysis was used to identify duplicates, redundancies, and non-clock sequences. Sequences were further verified by reciprocal BLAST (when the retrieved sequences served as a query in new searches) and/or by manual inspection of alignments. Several rounds of refined and reciprocal searches were performed in particular lineages and species to retrieve all available sequences. 2.2 Phylogenetic analyses Sequences were aligned using the MAFFT E-ins-i algorithm and phylogenetic trees were inferred using RAxML algorithm (Geneious Prime). For bHLH-PAS proteins, we used as reference Apis , Thrips , Drosophila , Danaus , and Halyomorpha sequences from the recently published dataset ( Tumova et al., 2024 ). To reconstruct the evolution of FBXL proteins, we utilized sequences from our early studies and a recent analysis of the cricket FBXL repertoire ( Takeuchi et al., 2023 ). As a reference of casein kinases, we used sequences from ( Thakkar et al., 2022 ), and for the remaining clock genes, we used the linden bug P. apterus ( Kotwica-Rolinska et al., 2022a ) and Drosophila . 2.3 Gene loss Whenever we failed to identify a clock gene in a particular species, we attempted to clarify whether this absence reflects insufficient sequencing (a very common situation especially in TSA), poor genome annotation, or if a gene loss is a likely explanation. As gene loss, we consider the situation when the gene is absent in all species of a monophyletic group for which well-sequenced genomes and transcriptomes are available2.4 Gene synteny To reconstruct gene synteny, we used the same approach as previously ( Smykal and Dolezel, 2023 ). First, we prospected genomic ( 6-4) photolyase locus in Homalodisca vitripennis (Auchenorrhyncha, Membracoidea superfamily) and genomic cry-DASH locus in Bemisia tabaci to identify protein-coding upstream and downstream syntenic genes. The syntenic genes’ coding sequences were then searched in TSA (Transcriptome Shotgun Assembly) databases of Nilaparvata lugens and Laodelphax striatellus (both Auchenorrhyncha, Fulgoroidea superfamily), or Trialeurodes vaporariorum (Sternorrhyncha, Aleyrodidae family), respectively, by BLAST (blastn, tblasn). The TSAs were then used as a query to identify genomic contigs/scaffolds with syntenic genes and to localize syntenic genes’ positions and the overall organization within the genomic regions. The number of protein-coding genes between syntenic genes for N. lugens and L. striatellus contigs is based on the original genome annotation. The cry-DASH -harboring region in Fig. 5 was BLASTED against GenBank to identify the source of the closest horizontal gene transfer donor. Syntenic gene IDs for Fig. 5 ( phr6-4 ) and Fig. 6 ( cry-DASH ) are in Supplementary Table S1 and Table S2, respectively. The resulting syntenies were drawn in CorelDRAW 6X (Alludo, Ottawa, Canada). 2.5 Gene isoforms and gene models To identify alternatively spliced isoforms, we used a similar approach as in ( Smykal et al., 2023 ), when we explored Pyrrhocoris apterus whole mRNA Oxford Nanopore Technology (ONT) reads. We used the longest m-cry and per coding sequences ( period : MW662133.1; m-cry : MW662132.1), blasted them against our custom-made P. apterus ONT brain transcriptomic databases and retrieved all mRNA reads. The reads were mapped to the in-house P. apterus genome using Minimap2 within Geneious Prime, allowing us to reconstruct the gene structure, determine the splice isoforms, and estimate their expression ratio. N. lugens and H. halys per and m-cry isoforms were primarily obtained from the annotated genomes, with some support from species-specific TSA databases. Protein domains (DNA_photolyase, FAD_binding_7, PAS, PAS_11, PeriodC) were predicted by using the PFAM database ( Mistry et al., 2021 ). The Nuclear Localization Sequences (NLS) were predicted using NLStradamus or Eukaryotic Linear Motif (ELM) resources ( Nguyen Ba et al., 2009 ; Kumar et al., 2022 ). The gene isoforms and gene model figures were drawn in CorelDRAW 6X. 3. Results 3.1. The ancestral clock First, we attempted to identify homologs of clock components known in insects ( Dolezel 2023 ; Tomioka and Matsumoto, 2015 ). Although this study focused on the clock gene inventory in the brown planthopper Nilaparvata lugens , we decided to interpret this inventory in the broader context of the entire group Hemiptera. As an outgroup, two thrips (Thysanoptera) species ( Frankliniella occidentalis and Thrips palmi ) and a body louse ( Pediculus humanus ) were used. As additional reference insects, we included the fruit fly Drosophila melanogaster , the black soldier fly Hermetia illucens , the red flour beetle Tribolium castaneum , the monarch butterfly Danaus plexippus , and the honeybee Apis mellifera ( Fig. 2 - 4 ). Download figure Open in new tab Fig. 2. Phylogeny of circadian clock proteins. (A) A phylogenetic tree illustrates the relatedness of insect PER to vertebrate PER1, PER2, and PER3. (B) Drosophila -type timeless (dTIM) and mammalian-type timeless (mTIM) are clearly distinguishable. (C) bHLH-Orange transcription factors. (D) Transcription factors with the basic leucine zipper domain (bZIP) include three clock components: Vrille (VRI), Par domain protein 1 (Pdp1), and Kayak (Kay)/Fos, which are well-separated from each other and related to ATF3 and JUN proteins. (E) Proteins with the basic helix-loop-helix PER-ARNT-SIM (bHLH-PAS) domain include three circadian clock components: CYCLE/BMAL, CLOCK, and TAIMAN (also known as NCoA or SRC). (F) Nuclear Hormone Receptors (NHR) include two circadian clock components: HR3 and E75. (G) Insect photolyases and cryptochromes form five clusters; however, the cry-DASH gene is found only in Bemisia . (H) Although the phylogeny of all F-box and leucine-rich repeat proteins (FBXL) is not easily reconstructed, the three proteins relevant to the circadian clock are well-defined. (I) Doubletime (DBT), Casein Kinase Iα (CKIα), and CKIγ form well-separated clades. Note duplications CKIα-like in Drosophila . NEMO and SHAGGY proteins were used as an outgroup. Consistent with early studies ( Yuan et al., 2007 ), the ancestral combination of clock components includes PER, dTIM, mCRY, and dCRY. This clock setup, characteristic to Lepidoptera, was identified in Thysanoptera, in which also FBXL3 protein was identified ( Kotwica-Rolinska et al., 2022a ) ( Fig. 1 ). FBXL3 and its close paralog FBXL21 participate in mammalian clock where these proteins bind to mCRYs ( Godinho et al., 2007 ; Siepka et al., 2007 ; Hirano et al., 2013 ). Thus, Thysanoptera seems to represent one of a few insect lineages with a complete set of circadian genes. However, the actual functional role of these genes has not been addressed. 3.2. Various reduction of clock components in outgroup species The additional, more distant outgroups than Thysanoptera, characterize the diversity of clock setups identified in insects. The body louse Pediculus humanus is an example of a species in which dCRY and JET have been lost. A similar loss was reported for A. mellifera and T. castaneum , however, more detailed species sampling revealed that all three losses of dCRY and JET are independent ( Kotwica-Rolinska et al., 2022a ). Furthermore, the honeybee (and all Hymenoptera) also lost dTIM and its clock setup, thus, resembling the mammalian one ( Rubin et al., 2006 ). The lepidopteran clock represented by the monarch butterfly Danaus plexippus contains the nearly complete clock gene toolkit when only FBXL3 is missing. The fruit fly Drosophila has lost mCRY and contains dCRY (in addition to (6-4) photolyase and CPD photolyase). Interestingly, Hermetia illucens , a dipteran species belonging to (infraorder) Stratiomyomorpha contains both genes. Thus, the loss of mCRY seems to be confined to a relatively narrow group of Cyclorrhapha (Muscomorpha). 3.3. Evolution of clock setup in Hemiptera Hemiptera is an assemblage of three groups, usually recognized as orders: Auchenorrhyncha, Sternorrhyncha, and Heteroptera. The phylogenetic relationships among them appear to be well resolved, thanks to modern phylogenomic approaches. The separation of Hemiptera from Thysanoptera is estimated to have occurred approximately 400 million years ago ( Johnson et al., 2018 ). As illustrated in Fig. 4 , the majority of circadian clock genes are present in all Hemiptera. An exception is d-cry , which has been lost in Halyomorpha (Pentatomomorpha) and Cimex (Cimicomorpha). This loss is accompanied by the loss of jetlag ( fbxl15 ). An independent loss of jetlag is observed in Aphidoidea (aphids and phylloxera species); however, these lineages possess d-tim . To clearly identify the m-cry gene, we have also annotated its related paralog, ( 6-4) photolyase ( phr6-4 ). This gene, which does not play a role in the circadian clock, has been lost independently in true bugs (Heteroptera) and Fulgoroidea ( Fig. 5 ). 3.4. Evolution of opsin genes and norpA Phylogenetic reconstruction of insect opsins revealed five well-separated groups ( Fig. 3A ). For simplicity, we have adopted the nomenclature used in a previous aphid-focused study ( Collantes-Alegre et al., 2018 ) to label these groups. The branches corresponding to Long-Wavelength Opsins (LWO), Rhodopsin 7 (Rh-7), Short-Wavelength Opsins (SWO), and Medium-Wavelength Opsins (MWO) contain only insect sequences. Download figure Open in new tab Fig. 3. (A) Insect opsins form five well-separated monophyletic groups described as follows: Long wavelength opsin (LWO); a monophyletic group comprising medium (MWO) and short (SWO) wavelength opsins; the rhodopsin 7 (Rh7) group; Arthropsin; and C-opsin. However, the group names reflect the names of proteins in representative species, whereas the actual wavelength sensitivities remain mostly unknown. A turquoise arrowhead highlights the SWO gene duplication observed in planthoppers (upper arrowhead) and aphids (lower arrowhead). (B) Phylogeny of Phospholipase C-21 (PLC-21C) and Phospholipase C-β (PLC-β), the latter known as No Receptor Potential A (NorpA). (C) Insect Quasimodo (QSM) phylogeny, where QSM from Basal hexapods and Crustacea ( Homarus and Daphnia ) served as outgroups. The distinction between the SWO and MWO groups in the phylogenetic tree is somewhat unclear and arbitrary (see Discussion for further details). To emphasize this ambiguity, we use quotation marks for these groups (“SWO” and “MWO”). Notably, three heteropteran groups have lost the “MWO” opsin ( Fig. 4 ). Interestingly, independent duplications of “SWO” have occurred in two of these groups, Fulgoroidea and Aphidoidea. Collantes-Alegre et al. (2018) suggested that an amino acid substitution from K90 to V90 in transmembrane domain 2 (TM2) could have caused a wavelength shift in one SWO in aphids. Such neofunctionalization might explain aphid sensitivity to medium wavelengths, even though no aphid opsin branches are found within the MWO cluster. Similarly, an independent duplication of the SWO gene in Fulgoroidea coincides with the loss of the MWO group. Furthermore, these SWO paralogs in Fulgoroidea are distinguished by a T-to-I substitution, located one amino acid downstream from the K90-to-V90 change observed in aphids (Supplementary Figure S1). Rh-7 opsins are present in all hemipteran species and in most reference insect species. They have been lost only in the honeybee and the red flour beetle. Among the five analyzed aphid species, four contain two Rh-7 genes. Closer examination suggests that this is the result of a specific duplication event ( Fig. 3A ). Additionally, the paralogous proteins differ at several amino acid positions (Supplementary Fig. S1). Human OPN4, which encodes melanopsin, branches between the previously described insect opsin groups (LWO, SWO, MWO, and Rh-7) and two additional groups: Arthropsins and C-opsins. Arthropsins have been lost in Thysanoptera, Psocodea, and holometabolous outgroups ( Fig. 4 ). Within Hemiptera, Arthropsins have only been lost in Halyomorpha (Pentatomomorpha). Ciliate opsins (C-opsins) branch as a sister group to Human OPN1 proteins (five paralogs), RHO, and OPN3. Two additional human protein sequences, OPN5 and peropsin, branch between C-opsins and Arthropsins. The final human protein, OPN4, which encodes melanopsin, branches between the Arthropsin/C-opsin group and the remaining insect opsins (LWO, MWO, SWO, and Rh-7; Fig. 3A ). In addition, several single-species gene duplications were identified, such as two Arthropsin genes in Homalodisca and two Rhodopsin 7 genes in Apolygus . However, these cases were not investigated further, and no attempt was made to determine whether they represent true gene duplications or errors in genome assembly. Two phospholipases, PLC-21C and NorpA, are relevant for circadian clock in Drosophila . Interestingly, the norpA gene has been duplicated in Thysanoptera, which is, to our knowledge, the only documented instance of norpA gene duplication in insects ( Fig. 3B ). 3.5. Unique cry-DASH in Bemisia CRY-DASH-type cryptochrome is absent in all insects except the silverleaf whitefly Bemisia tabaci . The phylogenetic analysis placed B. tabaci CRY-DASH, encoded by an intron-less gene, among plant and fungi CRY-DASH proteins and suggested a horizontal gene transfer (HGT) as a possible origin ( Kotwica-Rolinska et al., 2022a ). We inspected the B. tabaci cry-DASH locus and its flanking regions for a trace of donor, non-insect DNA. We first searched for seven protein-coding genes upstream and downstream of B. tabaci cry-DASH in the genome of related (but cry-DASH -less) whitefly Trialeurodes vaporariorum to estimate the size of the genomic region transferred from a host. Although spread over >47 Mbp, all seven genes were successfully identified and localized in the T. vaporariorum contig VMOF01000024.1. Roughly ∼ 130 kbp-long cry-DASH genomic region between syntenic genes was blasted against GenBank genome databases. The 33-kbp-long Solenum lycopersicum genomic contig was the only long positive hit, mapping ∼ 21 kbp downstream to the cry-DASH gene ( Fig. 6A ). The corresponding B. tabaci genomic sequence is ∼90% identical to S. lycopersicum contig with two suspicious short protein-coding genes unique to B. tabaci and a peculiar CANIN domain-containing gene ( Fig. 6B ). The S. lycopersicum genomic fragment is not mapped as a single uninterrupted fragment but as seven ≥700-bp-long fragments, suggesting further evolution of the locus in B. tabaci . Importantly, S. lycopersicum DASH is not localized within the 33-kbp DNA contig. Additionally, a comparison of coding sequences revealed only ∼40% identity between B. tabaci and S. lycopersicum CRY-DASH, in contrast to the ∼90% identity observed in downstream-located DNA. Previous analyses suggested that Bemisia CRY-DASH is related to plants or fungi; however, the phylogenetic relationship had not been reconstructed in detail. These discrepancies prompted us to expand the CRY-DASH sequence dataset and reanalyze the phylogenetic relationships ( Fig. 6C ). B. tabaci CRY-DASH clusters with fungal CRY-DASH proteins from several fungal lineages, particularly Ascomycota, and is clearly separated from the strictly plant branch. The presence of four plant CRY-DASH sequences identified in the Transcriptome Shotgun Assembly (TSA) databases within the ‘fungal’ cluster is highly suspicious and may represent cross-contamination of plant samples with fungal material ( Fig. 6C ). Although the cry-DASH gene is localized outside the donor region, the presence of non-insect DNA in its vicinity strongly supports horizontal gene transfer (HGT) as the most parsimonious explanation. 3.6. Novel clock gene isoforms identified in Hemiptera Gene presence or loss is not the only way to shape gene phenotypic output. We analyzed the splicing of mammalian-type cryptochrome ( m-cry ) and period ( per ) in several heteropteran species and focused on how it could affect the variability of the translated proteins. mCRY protein possesses two major domains, N-terminal DNA photolyase and C-terminal FAD binding 7 domain ( Fig. 7 ). In all species analyzed, N. lugens , P. apterus and H. halys , m-cry is a subject of alternative splicing, giving rise to the truncated isoforms. N. lugens m-cry has two predicted transcripts, each transcribed from an alternative exon. Transcript X1 encodes the full protein with complete DNA Photolyase and FAD binding 7 domains. The shorter isoform X2 transcription starts from non-coding (5’UTR) exon 4 and lacks ∼68% of the Photolyase domain. Similarly, H. halys isoform X4 starts from an alternative exon 4, which leads to the lack of ∼67% of the Photolyase domain. A remarkable splicing pattern was found in P. apterus and H. halys , where we found m-cry isoforms with retained intron 10, splitting the FAB binding 7 domain-encoding exons ( Fig. 7 ). Translation of retained-intron isoforms in both species leads to truncated mCRY proteins, with only ∼48% ( P. apterus ) and ∼22% ( H. halys ) fragment of the FAB binding 7 domain. The domain fragment in H. halys is below the threshold to be predicted as the FAD binding 7 domain by the PFAM database algorithm, and thus, the domain was not highlighted in Fig. 7 . Interestingly, the ratio of intron-retained/full m-cry mRNA reads reached ∼32% in P. apterus Oxford Nanopore Technology brain transcriptomes. In P. apterus , C and D per isoforms are likely destroyed via NMD if translated from the first canonical AUG (isoforms C1 and D1) or translated into PER isoforms C2 and D2 if (downstream) alternative translation start site is preferred ( Fig. 7 ). Whether initial codon skipping happens in vivo and gives rise to C2 and D2 isoforms is to be determined. Download figure Open in new tab Fig. 4. Inventory of circadian clock setup and light input components. Representative insect species are shown with gene presence (full circle) or absence (empty circle) indicated. Lineage-specific losses are highlighted with magenta rectangles, while lineage-specific duplications are highlighted with turquoise rectangles. Numbers represent the presence of multiple paralogs within a single taxon. Abbreviations are as follows: FULGOR. – Fulgoroidea, MEMBR. – Membracoidea, PSYLLO. –Psylloidea, THYSAN. –Thysanoptera, PSOCOD. –Psocodea, HYMENOP. –Hymenoptera, COLEOP. –Coleoptera, LEPIDOP. –Lepidoptera. Download figure Open in new tab Fig. 5. Gene synteny supports the loss of (6-4)-photolyase ( phr 6-4 ) in Nilaparvata lugens and other Fulgoroidea. Laodelphax striatellus and N. lugens (both Fulgoroidea) have lost phr6-4 , which remains present in the sister family Membracoidea, as represented by Homalodisca vitripennis . The phr6-4 upstream and downstream syntenic genes from H. vitripennis are scattered across a single contig or scaffold in the genomes of N. lugens and L. striatellus . Syntenic genes are color-coded for easier tracking. Numbers between genes in N. lugens and H. vitripennis indicate the number of protein-coding genes separating the respective syntenic genes. Note the significant reorganization of the presented genes within the contigs. Download figure Open in new tab Fig. 6. Bemisia tabaci cryptochrome-DASH locus analysis supports horizontal gene transfer. (A) Seven protein-coding genes located upstream and downstream of the B. tabaci cryptochrome-DASH were also found in the related whitefly Trialeurodes vaporariorum ; however, the gene order is significantly rearranged. These syntenic genes are color-coded for easier tracking (note that each sequence is presented at a different scale). Interestingly, a BLAST search of the B. tabaci cry-DASH locus DNA sequence against GenBank genome databases revealed ∼90% identity between a 33-kbp genomic fragment of the tomato plant ( Solanum lycopersicum ; JALGYV010003334.1) and a ∼40-kbp segment of the B. tabaci genomic sequence (NW_017548003.1) located downstream of the cry-DASH gene. (B) Details of three genes predicted in the locus downstream of cry-DASH . Note that the protein-coding sequence constitutes only a small portion of each predicted hypothetical gene. (C) Phylogenetic analysis of CRY-DASH proteins. Animal CRY-DASH proteins served as an outgroup (top, Animalia). One cluster contains only CRY-DASH from green plants (green background, middle part). The bottom branch includes several representatives of fungal lineages (Chytridiomycota, Mucoromycota, Zoopagomycota, Basidiomycota, and Ascomycota), highlighted with different background colors. The Bemisia cry-DASH branches within Ascomycota . Four plant “CRY-DASH” protein sequences branching within fungi were identified in the Transcriptome Shotgun Assembly (TSA) databases. These sequences are highly suspicious and are likely the result of cross-contamination of the plant sample with fungi. Download figure Open in new tab Fig. 7. mCRY splicing isoforms. The m-cry gene of N. lugens , P. apterus , and H. halys is subject to alternative splicing, which generates some truncated mCRY protein isoforms. In N. lugens , transcription from an alternative exon 4 produces an mCRY isoform with a significantly shorter DNA photolyase domain. Retained intron 10 in P. apterus and H. halys introduces a premature stop codon, truncating the coding sequence of the mCRY FAD-binding 7 domain. These shorter mCRY proteins are either non-functional or targeted for transcript suppression via nonsense-mediated mRNA decay. All exons are shown to scale and numbered in the 5’ → 3’ direction, with letters depicting exon sub-variants. The black arrowhead marks the positions of P. apterus m-cry mutants (Kotwica-Rolinska et al., 2022). A complex splicing pattern was found in the 5’ region of the period gene in the analyzed species, giving rise to PER variants differing in the N-terminal part. Reference D. melanogaster period is not alternatively spliced in this gene region. The only translated PER protein isoform possesses an N-terminal nuclear localization signal (NLS, Chang and Reppert 2003 ), two PAS domains, mid-protein NLS ( Fig. 8 ), and several phosphocluster regions, which are phosphorylated to ensure PER stability, inhibitory potency and modulation of temperature compensation phenotype, forming thus a phospho-switch to balance diverse PER functions throughout the day-night cycle ( Joshi et al., 2022 ). One of the phosphoclusters, the N-terminal phosphodegron, is responsible for PER degradation in the proteasome ( Chiu et al., 2008 ; Kivimäe et al., 2008 ). It includes Serine 44,45 and 47 (numbering according to D. melanogaster PER, Chiu et al., 2008 ). S47 is phosphorylated by DBT and together with nearby phosphorylated sites, phospho-S47 generates a high-affinity atypical SLIMB-binding site ( Chiu et al., 2008 ). The phosphodegron with flanking sequences is conserved in full PER protein isoforms in analyzed species, PER isoform X1 in N. lugens , isoform A in P. apterus and isoform X1 in H. halys ( Fig. 8B ). Unlike D. melanogaster , analyzed heteropteran PERs contain an additional C-terminal Period circadian-like domain (PeriodC domain, PFAM ID: IPR022728) ( Fig. 8A ). The full PER represents only a subset of possible splice variants in analyzed hemipteran species, with several gradually truncated PERs. In N. lugens and H. halys , transcription from alternative exons 9 and 7 leads to an expression of X19 and X8 isoforms, respectively, missing the phosphodegron and N-terminal NLS. Most intriguing per isoforms, generated by the alternative splicing, were detected in P. apterus and H. halys . Transcription of P. apterus per isoform B from exon 2 and H. halys per isoforms X6 (and X7) from exon 3 alternative transcription start site reveals the first initiation codon (AUG) in exon 4 but in reading frame 2 (depicted by a red line in Fig. 8A ). The same region is in full-PER isoforms read in frame 1. In both species, translation starts with exon 4 and the translation in reading frame 2 continues through exon 6 ( P. apterus ) or 5 ( H. halys ). The absence of exon 7 ( P. apterus ) or exon 6 ( H. halys ) in the ‘alternative’ B and X6/X7 isoforms, respectively, brings the rest of the open reading frame into line with reading frame 1, thus avoiding premature stop codon and enabling translation of the downstream canonical PER sequence and domains (PAS, mid-protein NLS, PeriodC). Remarkably, the exons with dual/overlapping reading frames encode NLS in both frames (!), although the exact AA composition differs. P. apterus full-length mRNA Oxford Nanopore Technology (ONT) reads further revealing the presence of minor versions of transcripts encoding isoforms A and B, in Fig. 8 presented as isoforms C and D. Those isoforms either lack (isoform C) or contain (isoform D) frame-adjusting exon 7, thus decoupling the frames. Download figure Open in new tab Fig. 8. PERIOD N-terminal isoforms are generated by alternative splicing and dual-reading-frame translation of specific exons. (A) Full-length PER proteins are translated in reading frame 1 (brown line) and possess conserved 32-33-amino-acid-long N-terminal PER region (yellow line). A gradient of N-terminally truncated PER is generated by alternative splicing in N. lugens , P. apterus and H. halys , but not Drosophila melanogaster . Some isoforms lack conserved N-terminal PER region, some lack N-terminal nuclear localization signal (NLS). N. lugens per isoforms are generated by transcription from alternative transcriptional start sites. Transcription of P. apterus per isoform B and H. halys per isoforms X6 and X7 from alternative transcription start sites leads to translation in reading frame 2 (red line). The absence of exon 7 ( P. apterus ) or exon 6 ( H. halys ) in the isoforms changes the reading frame back to frame 1, enabling the presence of the canonical PER domains (PAS, mid-protein NLS, PeriodC). Interestingly, dual/overlapping reading frames exons encode NLS in both frames. P. apterus per isoform C2 is likely destroyed via nonsense-mediated mRNA decay if translated from the canonical AUG or results into isoform C1 if (downstream) alternative translation start site is preferred. All exons are numbered in a 5’->3’ direction and presented in scale, except H. halys exon 7, which was shortened for space reasons. The black arrowhead depicts the positions of P. apterus per mutants ( Kotwica-Rolinska et al., 2022a ). See Methods and Materials for a detailed description of the protein domain and motifs’ prediction. (B) The alignment of conserved N-terminal PER region. The black rectangle highlights Serine 47 (S47), targeted in Drosophila melanogaster by DBT and upon phosphorylation targeting PER for degradation ( Chiu et al., 2008 ). The yellow line depicts the conserved N-terminal PER region and the blue curved line predicted NLS. 4. Discussion This study focused on circadian clock genes and opsins in Hemiptera. In principle, this is a gene inventory aimed at shedding light on the evolution of this time-measuring molecular device. While some earlier studies focused on individual species, such as the bean bug Riptortus pedestris ( Ikeno et al., 2008 ) or the pea aphid Acyrthosiphon pisum ( Cortés et al., 2010 ; Barberà et al., 2017 ), systematic comparisons, to our knowledge, have not been provided. Our earlier publication ( Kotwica-Rolinska et al., 2022a ) addressed the function of the majority of clock genes in the linden bug Pyrrhocoris apterus ; however, the comparative part was limited to PER, dTIM, mTIM, cryptochromes, JET, and FBXL3 proteins. In addition to Hemiptera, our current description includes several reference/outgroup insect lineages. Thus, we can further expand the concept of what the ancestral type of insect circadian clock consisted of. The combination of mCRY and dCRY, typical of the lepidopteran clock, is further complemented by FBXL3 in Thysanoptera, suggesting that, at least at present, thrips possess the most complete circadian gene setup among insects. The reference insect species illustrate the diversity of clock setups in insects, including the notable loss of dCRY, (6-4) photolyase, and JET. In the case of Apis , the clock setup is further reduced by the loss of dTIM. The comparison of Drosophila and Hermetia , both members of Diptera, illustrates the mCRY loss unique to Cyclorrhapha. Although some important differences in the mechanism of the circadian clock exist, such as the involvement of mCRY in numerous species contrasting with Drosophila ( Ikeno et al., 2011a ; Zhang et al., 2017 ; Werckenthin et al., 2020 ) and the altered function of dTIM ( Kamae and Tomioka, 2012 ; Kotwica-Rolinska et al., 2022a ), it is still feasible to extrapolate clock properties based on the Drosophila mechanisms. For example, the loss of dCRY and JET in P. apterus fits with the lower sensitivity to light for entrainment ( Kaniewska et al., 2020 ). In aphids, dCRY is unexpectedly stable upon constant light illumination ( Colizzi et al., 2021 ). This stability can be explained by the loss of JET and further correlates with the accelerated evolution of dTIM in aphids and phylloxera ( Bullo et al., 2024 ). Whether dCRY participates in the light entrainment of the circadian clock or plays some role in the photoperiodic timer in aphids, remains elusive. However, the light input to the clock is rather complex when parallel dCRY-independent pathways have been reported. In some cases, it is not clear whether they represent a conserved mechanism or are mere species-specific idiosyncrasies. For example, the role of QSM in light responsiveness has not been explored beyond Drosophila melanogaster . In the case of opsins, their role in the circadian clock is well-established in several insect species. However, the presence of multiple opsin paralogs makes their research technically challenging. For example, seven opsin genes are recognized in Drosophila . Thus, despite the remarkable genetic tools available to this model organism, even combining all available mutations into one strain becomes a non-trivial task. For example, the role of Rh-7 is still being uncovered ( Senthilan and Helfrich-Förster 2016 ). A second research challenge is the spectral overlap in the sensitivity of several opsins. Importantly, the spectral sensitivity has been experimentally determined for only a minimal number of opsins, whereas the properties of many others remain elusive. Therefore, the definition of particular groups in this study should be understood based on their evolutionary position rather than their actual light-detection properties. Notably, some experimental evidence highlights specific residues that are key for determining wavelength sensitivity. When these residues are mutated, spectral shifts occur, as suggested for one of Aphidoidea (and possibly for Fulgoroidea) SWO paralogs ( Collantes-Alegre et al., 2018 ). Such light-sensing adaptability would contribute to physiological plasticity and potentially be a simple solution compensating for MWO gene loss. In addition to gene losses, we identified a unique horizontal gene transfer in B. tabaci , where a fungal cry-DASH had been inserted into the insect genome. The presence of an exogenous DNA in the vicinity of the cry-DASH locus in the B. tabaci genome supports HGT as a feasible origin of cry-DASH in only one insect species. Our analysis shows that the Solanum lycopersicum genomic fragment is mapped in several neighboring regions, suggesting ongoing whitefly-specific evolution of the locus. Although S. lycopersicum seems to be the donor of the HGT sequence, we cannot rule out the possibility that the true donor species has not been sequenced yet and S. lycopersicum genomic fragment is just the closest species present in the GenBank. Phylogenetic analysis of CRY-DASH proteins questioned the plant origin of B. tabaci CRY-DASH, placing it into a ‘fungal-specific’ cluster. Indeed, B. tabaci CRY-DASH is more related to fungal than plant CRY-DASH proteins, including CRY-DASH from S. lycopersicum genome. Whether the physical vicinity of B. tabaci cry-DASH locus and exogenous DNA represents a single or two independent HGT events is unclear. The gene synteny comparison with the related Trialeurodes vaporariorum whitefly genome underpins major genomic rearrangements, further masking the origin of the cry-DASH locus. In any case, B. tabaci is a polyphagous species with ∼500 host plants in its repertoire, thus facing many challenges when feeding upon different hosts. Over 49 genes were likely obtained from plant hosts via HGT by B. tabaci ( Lapadula et al., 2020 ; Xia et al., 2021 ; Gilbert and Maumus 2022 ), many linked to insect-plant interactions. The HGT can provide an alternative for gaining new capabilities through classical gene duplication and evolution. Although the cry-DASH in B. tabaci is transcribed, whether possessing this gene constitutes an advantage remains to be determined. The transcriptome analysis revealed a remarkable dual reading frame in P. apterus and H. halys period transcripts. The Serine-rich phosphodegron is well conserved in full-length PER proteins in all four analyzed species, with P. apterus PER lacking Ser47 (replaced by Ala50). At least one isoform lacking PER phosphodegron is annotated in N. lugens , P. apterus , and H. halys , probably omitting DBT– and SLIMB-specific regulation in the N-terminal region ( Fig. 8A ), however, specific point mutations in D. melanogaster PER phosphodegron keep downstream parts of the protein still accessible to DBT and SLIMB regulation ( Chiu et al., 2008 ). Unlike Drosophila , where PER heterodimerizes with and is stabilized by dTIM, all analyzed species possess mCRY ( Fig. 3 and Kotwica-Rolinska et al., 2022a), a typical mammalian PER partner. We can speculate if phosphodegron-less PERs represent less SLIMB-sensitive isoforms or denote the variability of PER heterodimerization with dTIM and mCRY. The presence of phosphodegron-less PER isoforms in P. apterus and H. halys suggests that the lack of phosphodegron outweighs the AA variability in N-terminal NLS, encoded by the dual-frame exons, and a necessity to bring those exons in frame with the rest of the PER protein to contain PAS and other downstream domains. Manipulation of N-terminal and mid-PER NLS sequences in D. melanogaster PER suggested the mid-PER NLS is the main nuclear localization signal, however, N-terminal NLS still participates in PER nuclear import ( Chang and Reppert, 2003 ). Thus, an ‘alternative’ NLS with a different AA composition might still be sufficient to complement mid-PER NLS. Alternative splicing serves as yet another layer for controlling clock gene expression. For instance, retention of an intron in the central part of the d-tim gene in Drosophila results in transcripts that cannot encode a stable dTIM protein ( Martin Anduaga et al., 2019 ; Foley et al., 2019 ; Shakhmantsir et al., 2018 ). Furthermore, these mRNAs contain premature stop codons and are therefore likely destroyed by NMD. The high frequency of intron retention in the m-cry transcripts of P. apterus invites speculation that a similar regulatory mechanism might be at play. It will be interesting to determine whether similar intron retention occurs in other species. Since the stability of RNA-RNA interactions is temperature-dependent, alternative splicing is often influenced by temperature. Indeed, the temperature-dependent splicing of d-tim in drosophilid flies likely serves as a regulatory mechanism. Whether m-cry splicing is affected by temperature remains unknown, as does its potential regulatory role in the circadian clock. However, because the mCRY protein is a key clock component in numerous species, including P. apterus , significant regulation of its levels could, at least theoretically, have a profound impact on clock function. What is responsible for the observed genetic flexibility of clock setups? As these time-measuring devices are supposed to perform a conserved task, which is to keep ticking with a period of 24 hours, one would assume that the genetic components should be very conserved. While the general mechanism of the transcriptional/translational feedback loop is found in both deuterostomes (including mice, humans, and zebrafish) and protostomes (insects, mollusks, and annelids), the participating players are conserved only partially. Either gene losses or a high degree of sequence variability in the protein sequences are observed. One possible mechanism allowing for gene loss and rapid evolution of some components is the functional redundancy of the system. For example, the light input to the clock is achieved by several pathways; thus, alteration of one of them does not result in complete “clock blindness.” Therefore, losing one of these pathways only impacts sensitivity, but the organism still perceives particular stimuli. This change in sensitivity might even be beneficial. For example, insects have colonized various environments that differ in photoperiod (day-to-night length). In the most extreme latitudes, nearly constant light is available during summer. Under such conditions, lower light sensitivity might be an advantage. Indeed, this is the case for Drosophila ezoana , which is partially rhythmic even under constant light conditions. Furthermore, this and some other species from high latitudes, such as Chymomyza costata , are not rhythmic in constant darkness ( Bertolini et al., 2019 ). In the latter species, a unique recessive mutation in the d-tim gene is associated with a malfunction in photoperiodic timing and impacts the expression of additional clock genes ( Pavelka et al., 2003 ; Stehlik et al., 2008 ; Kobelkova et al., 2010 ). The possible connection between photoperiodic timers and the circadian clock has been long discussed ( Kostal 2011 ; Dolezel 2015 ; Saunders 2005 , 2010 ; Bradshaw and Holzapfel, 2010 ). The recruitment of circadian clock components by the photoperiodic timer seems to be a logical explanation for establishing a relatively complex device measuring photoperiod from already available “tools”. Indeed, this view is further supported by the participation of several clock genes and circadian neuropeptide PDF in the photoperiodic timer in several species of Hemiptera ( Ikeno et al., 2010 ; Ikeno et al., 2011b , 2011c , 2013 ; Urbanova et al., 2016 ; Kotwica-Rolinska et al., 2017 , 2022b ; Kaniewska et al., 2024 ). However, although the evidence is impressive, the actual involvement of the entire circadian clock machinery in the photoperiodic timer has not been explored and the connection between the two devices is more complicated. One important critique is that only some components might have been recruited ( Emerson et al., 2009 ). Gene pleiotropy, such as when circadian clock components in the insect gut interact in a non-canonical way, suggests that the connection between the circadian clock and seasonality is indeed more complex ( Bajgar et al., 2013a , 2013b ; Dolezel et al., 2007 ). The circadian clock genes also define the activity phase, and this role might have shaped their evolution. In this regard, notable geographic variability in the free-running period has been reported for the linden bug P. apterus , for the wasp Nasonia vitripennis , and in the eclosion rhythm of Drosophila littoralis and D. subobscura ( Pivarciova et al., 2016 ; Paolucci et al., 2019 ; Lankinen, 1986 ; Lankinen, 1993 ). Another key role of the circadian clock is related to navigation during long migrations. The position of the Sun serves as a suitable orientation cue as long as the organism can compensate for its positional change during the day. The time-compensated sun compass is intensively studied in the iconic monarch butterfly, Danaus plexippus , in which the ability to detect the magnetic field contributes to migration either by fine-tuning the navigation system or serving as a backup mechanism ( Merlin et al., 2020 ). Moreover, the role of these components is further complicated by the participation of cryptochromes in the ability to detect the magnetic field and relay this information. The interference of the magnetic field with the effects of light has been well documented in Drosophila and the German cockroach Blattella germanica ( Fedele et al., 2014a , 2014b ; Yoshii et al., 2009 ). In the case of Drosophila , the response requires dCRY. The strongest phenotypes pointing to the role of dCRY in directional magnetoreception have been observed in the monarch butterfly ( Wan et al., 2021 ), although, in some species, mCRY has been suggested ( Bazalova et al., 2016 ; Netusil et al., 2021 ). Interestingly, a change in geomagnetic field intensity alters migration-associated traits in Nilaparvata ( Wan et al., 2020 ). The complex involvement of the circadian clock genes in various time-measuring devices, navigation, magnetoreception, and seasonality provides interesting scientific problems but may also be connected to applied research. Numerous hemipteran species are important pests and understanding the genetic basis of their daily activity preference, seasonal calendar, and navigation can be practical. Similarly, our analysis points to the importance of alternative splicing isoforms, such as the non-canonical PER variants and intron retention in m-cry . In both cases, and many others found in the future, it will also be interesting to explore whether the alternative splicing is affected by temperature. Perhaps, some alternative splicing events even participate in temperature compensation mechanisms or contribute to the regulation of temperature-dependent daily activity patterns. Therefore, it is important to identify and annotate all predicted gene isoforms when dealing with genomic or transcriptomic data. As recently shown by Chikhaoui et al. (2024) , the loss of PER proteins in the liver of mPer1 / mPer2 double mutant mice seems to directly alter alternative splicing likely through the mis-localization of serine/arginine-rich splicing factors (SRSF) within the nucleus. Thus, alternative splicing might represent still rather undervalued level of clock regulatory mechanism. While the in silico predictions point to important changes and irregularities in the clock setup, functional analysis is key for identifying underlying mechanisms. In this regard, stable genomic modifications are the most powerful approach to entering hemipteran research including challenging groups such as aphids ( Xue et al., 2018 ; Kotwica-Rolinska et al., 2019 ; Reding and Pick, 2020 ; Le Trionnaire et al., 2019 ; de Souza Pacheco et al., 2022 ). To make gene editing accessible to a broader range of non-model species, a more versatile means of Cas9/gRNA delivery are emerging ( Mocchetti et al., 2024 ; Shirai et al., 2022 ) and less harmful DNA-editing enzymes are tested ( Thakkar et al., 2023 ; Doll et al., 2023 ; Li et al., 2018 ). Therefore, we can assume that some significant progress will be made in understanding the circadian clock genes in either interesting biological problems or in agronomically important pest species. Author contribution Conceptualization: D.D., V.S., Data curation: V.S., H.T., D.D., Formal analysis: D.D., V.S., H.T., Funding acquisition: D.D., Investigation: D.D., V.S., H.T., Methodology: D.D., V.S., Resources: D.D., V.S., Supervision: D.D., V.S., Validation: D.D., V.S., Visualization: D.D., V.S., Writing – original draft: D.D., V.S., Writing – review and editing: D.D., V.S. Funding This work was supported by the Czech Science Foundation (GACR, 22-10088S) to D.D. Appendix A. Supplementary data Fig. S1A. Protein alignment of selected opsins from the Rh7, SWO, and MWO clades. Fig. S2 Phylogenetic analysis of CRY-DASH proteins. Table S1. Synteny in 6-4 photolyase locus Table S2. Cryptochrome-DASH synteny View this table: View inline View popup Download powerpoint Table 1. An enhanced list of issues to focus on if studying clock genes in heteropteran insects. References 1. ↵ Allada , R. , White , N.E. , So , W.V. , Hall , J.C. , Rosbash , M. , 1998 . A mutant Drosophila homolog of mammalian Clock disrupts circadian rhythms and transcription of period and timeless . Cell 93 , 791 – 804 . doi: 10.1016/s0092-8674(00)81440-3 OpenUrl CrossRef PubMed Web of Science 2. ↵ Bai , J. , Uehara , Y. , Montell , D.J. , 2000 . Regulation of invasive cell behavior by taiman, a Drosophila protein related to AIB1, a steroid receptor coactivator amplified in breast cancer . 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